Imaging and Quantifying the Morphology and Nanoelectrical

Nov 29, 2010 - School of Civil and Environmental Engineering, Georgia Institute of Technology, Atlanta, Georgia, .... Cytotechnology 2017 69 (2), 245-...
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J. Phys. Chem. C 2011, 115, 599–606

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Imaging and Quantifying the Morphology and Nanoelectrical Properties of Quantum Dot Nanoparticles Interacting with DNA Wen Zhang, Ying Yao, and Yongsheng Chen* School of CiVil and EnVironmental Engineering, Georgia Institute of Technology, Atlanta, Georgia, 30332 ReceiVed: August 13, 2010; ReVised Manuscript ReceiVed: NoVember 1, 2010

Rapid detection and imaging the interactions between quantum dots (QDs) and DNA are of interest for research on applications as well as for determining their potential cytotoxicity. This paper introduces Kelvin force microscopy (KFM), an electric mode of atomic force microscopy, as a technique to examine the binding of QDs with DNA in vitro and in vivo. KFM provides information about the nanoelectrical properties of QDs and DNA that is complementary to the topography and phase images that conventional AFM provides. With this unique function, KFM demonstrated its ability in determining the morphological and electrical changes in DNA after exposure to QDs as well as to distinguishing individual QDs from DNA matrices. Our results indicated that the nonspecific binding with QDs led to transformation of polymeric DNA into pearl-like spheres. In vivo experiments showed that QDs could permeate into E. coli cells and bind with genomic DNA. To the best of our knowledge, this is the first successful demonstration of KFM to detect single QD-DNA binding. KFM potentially can be used in characterization of nanomaterials and their interfacial interactions with biomolecular matrices. 1. Introduction Quantum dots (QDs) have similar sizes as proteins and other biomacromolecules and exhibit versatile unique properties.1-3 As a result, QDs have a wide range of applications such as medical diagnostics,4 drug delivery,5 DNA-based nanosensors,6,7 and solar energy conversion.8 With their increasing production and applications, QDs are more likely to interact with the environment and the human body, which raises concerns about their potential environmental and health impacts.9-12 Owing to its ultrasmall size, a single QD can interact with a biomacromolecule directly. Recent studies have provided compelling evidence on potential cytotoxicity caused by QDs.12,13 Our previous work showed that QDs (CdTe) coated with hydrophilic thioglycolate capping ligands that could remain in finished water after conventional water treatment processes14 disrupted the epithelial monolayer of human intestinal cells (Caco-2) and caused cell death at 0.1 mg/L of QDs.15 Other studies demonstrate similar exposure impacts such as genotoxic effects on human cells11 as well as bactericidal,16 and pathogencidal effects.10 As pointed out previously, these biological effects may be significantly governed by the nanobio interfacial interactions between QDs and cells.17 However, such interactions with QDs involved are still poorly understood, especially at molecular genetic levels. Despite advancements in the application of QDs in gene therapy18 and DNA sensing,19 precise control of such bioconjugation has remained elusive.20 Therefore, biological interactions with QDs are gaining considerable attention because uncontrolled interactions such as nonspecific adsorption, may lead to undesirable effects (e.g., genotoxicity).21,22 Obviously, deoxyribonucleic acid (DNA) is a potential target of these unintentional attacks of ultrasmall QDs. Because DNA carries genetic information and regulates the synthesis of enzymes and * To whom correspondence should be addressed. E-mail: [email protected]. Phone: (+1) 404-894-3089. Fax: (+1) 404894-2278.

other proteins,23 damages to it can lead to disease initiation and progression.24,25 Recent studies indicated that QDs could cause DNA damages26 and nick supercoiled DNA.27 However, these studies provided no visualization of QD-DNA interactions. DNA interaction with carbon nanoparticles (CNPs) were examined with atomic force microscopy (AFM) and DNA aggregation was observed.28 This study employed only conventional AFM to investigate the morphological impacts on DNA, but local identification of individual CNP and DNA molecules were not achieved. The binding mechanisms are therefore still largely unknown. Furthermore, hydrophilic QDs may have dramatically different interactions with DNA than hydrophobic CNPs, which is worth exploration. Currently, few methods are available for detection of interacting entities at the nanoscale and for quantifying and characterizing exposure to nanomaterials in biological matrices. Such tools are needed to reveal biophysicochemical interactions at the nanobio interface and to understand how interfacial properties affect the biological consequences upon exposure.24 In this regard, Kelvin force microscopy (KFM), an electric mode of atomic force microscopy (AFM), was explored in this study to examine the nanoelectrical properties and to map the surface heterogeneities of QDs and DNA.29 KFM provides information about nanoelectrical properties (primarily referred to as surface potential at nanoscale in this study) that is complementary to the topography and phase images generated by standard AFM,30-32 which can serve locally to identify and differentiate individual NPs. KFM is based on the Kelvin method except that electrostatic forces acting on the cantilever are measured instead of AC currents. A DC power supply is used to bias the probe, and the AC source introduces an alternating force component in the cantilever that depends on the intrinsic surface potentials of the target materials. Electrostatic forces between the probe and regions in the sample arise due to differences in local mechanical and electromagnetic properties,33 such as surface charge, doping level, and dielectric constant. A DC

10.1021/jp107676h  2011 American Chemical Society Published on Web 11/29/2010

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voltage is applied to nullify this difference, and the surface potential can thus be quantified.34,35 QDs (CdSe/ZnS, core/shell), double-stranded DNA homopolymers [poly(dA) · poly(dT) and poly(dG) · poly(dC)], and genomic DNA extracted from E. coli cells were examined by KFM to resolve their original morphology and electrical properties as well as changes in those properties after in vitro and in vivo binding experiments. The overall goal of this research is to provide insight into the interactions of genomes with QDs using KFM as a novel imaging and quantification tool. 2. Experimental Section 2.1. Quantum Dots. Water-soluble QDs were purchased from Ocean NanoTech, LLC. The QDs were CdSe/ZnS core/ shell coated with hydrophilic capping ligands (carboxylic acid). This type of QDs is photostable due to the shielding effect from the shell and thus no photoexcitation effects are considered.12,36 The QD stock suspension had a concentration of 8 µM and the pH was between 10 and 11. The vendor-reported size of the QDs is 18.4 ( 4.2 nm. The concentration of Cd in the QDs was further analyzed by an inductively coupled plasma-mass spectrometry (ICP-MS, DRC Elan II, PerkinElmer, USA), as previously reported12,14,15 The QD sample was digested with high-purity HNO3 (67-70%, w/w) at 125 °C for 25 min based on the ASTM the standard method.37 2.2. Homopolymer DNA. Powder forms of DNA homopolydeoxynucleotides, poly(dA) · poly(dT) and poly(dG) · poly(dC), were purchased from Sigma-Aldrich (catalog nos. P9764-10UN and P3136-10UN, respectively). The two homopolymers as obtained were suspended in 1 mL of TE buffer (pH 8.0, 10 mM Tris-HCl, 1.0 mM EDTA) and due to the lower solubility of the poly(dG) · poly(dC) polymer that suspension was heated to 50-55 °C for 30 min with occasional mixing by inversion. The stock solution (500 ng/µL) was diluted with TE buffer and stored at 4 °C. Before mixing with QDs, the two homopolymer suspensions were heated in a hot water bath at 95 °C for approximately 30 min to effect full dissociation. 2.3. In Vitro DNA Binding with QDs. QDs were added into a 1.5 mL centrifuge tube at a final concentration of 8 nM with DI water, and then the poly(dA) · poly(dT) DNA was dispersed into the tube at a final concentration of 10 ng/µL. The tube was stirred with a vortex for 30 s and then incubated at 37 °C for 1 h. The same operation was applied to poly(dG) · poly(dC) DNA. 2.4. Culture of E. coli Cells and in Vivo DNA Binding with QDs. E. coli K-12 cells (strain D21) were purchased from the E. coli Genetic Stock Center (Department of Biology, Yale University, New Haven, CT) and were cultured in standard Luria-Bertani (LB) medium at 37 °C overnight. To determine the cell growth, OD (optical density) was measured at a wavelength of 600 nm with a spectrophotometer (Beckman, DU7700). In the exposure experiments, 200 µL of cell culture were harvested at the midexponential growth stage (OD600 of this culture was 1-1.5, and the cell density was approximately 2 × 109 cells/ml) and then inoculated into 5 mL fresh LB medium containing different concentrations of QDs (0, 30, 90, or 200 ppb based on [Cd] concentration). This secondary culture was incubated at 37 °C for 12 h with vigorous shaking at 200 rpm. OD600 was monitored to study the exposure effect of QDs on the growth of E. coli cells. Total DNA was extracted from the cultured E. coli cells with and without exposure to 200 ppb QDs using the Qiagen DNeasy

Zhang et al. Blood and Tissue Kit. Briefly, a pellet of 1 mL of cell suspension with a concentration of approximately 2 × 109 CFU/ml was placed in a 1.5 mL centrifuge tube and centrifuged for 10 min at 5000×g. The supernatant was discarded and the pellet resuspended in 180 µL of ATL buffer (see the Quagen manual for more introductions to this buffer). The total DNA was extracted following the protocol of the Qiagen kit. DNA yield was determined by measuring its absorbance at 260 nm with a Beckman spectrophotometer (an A260 value of 1 with a 1 cm detection path corresponds to 50 µg DNA/ml). The A260/A280 ratio was used to evaluate the protein contamination of the DNA (normally within 1.8-2.0). 2.5. KFM Study of the DNA Binding with QDs. The KFM was operated on an Agilent 5500 AFM (Molecular Imaging) equipped with MAC III unit, which has three lock-in amplifiers (LIA) enabling multifrequency measurements. LIA 1 was used for topography imaging in the intermittent contact mode, which was performed at the first flexural resonance of the probes ωmech and LIA 2 was used for KFM, providing AC and DC voltages to the probe and detecting the electrostatic response either from the photodetector (AM-AM) directly or from the LIA 1 employed for the topography servo (AM-FM). Pt-coated silicon cantilever probes (Olympus AC240TM, Japan) with a force constant of approximately 2-5 N/m and a nominal resonance frequency of 70 kHz were used and operated with an offset of 0.6 V below the measured resonance frequency. An AC bias voltage of 2 V at a frequency of 70 kHz was applied between the probe and sample and a DC bias was set at a frequency of 5 kHz with the bias voltage provided from the LIA 2 servo. In KFM, the whole microscope was fully contained in an environmental chamber that was used to control ambient pressure, temperature (25 ( 2 °C), and humidity (approximately 35%) measured by VWR humidity/temperature thermometer. The stock solution (8 µM) of QDs was first diluted by 1000 times and then 0.5 µL of the diluted solution was deposited on a clean silicon wafer purchased from Sigma-Aldrich that had been cleaved to a small piece of approximately 3 mm × 8 mm and air-dried for 1 min. The silicon wafer was finally fixed on a small piece (1 cm × 1 cm) of conductive double-sided tape (Ted Pella), which was placed on a grounded microscope stage. Topography, phase, and surface potential images were obtained simultaneously. To examine the homopolymer DNA and E. coli DNA before and after exposure to QDs, the original DNA solution was diluted to a concentration of approximately 10 ng/µL and 0.5 µL was taken and dried on the silicon wafer substrate. After air for 1 min, the sample was ready for KFM study. For the DNA mixed with QDs, 0.5 µL was directly taken and used for KFM study. 3. Results and Discussion 3.1. Characterizations of QDs. The obtained topography and morphology of QDs indicated that they were spherical in shape and had a relatively uniform size distribution (see the white dots in Figure 1a). According to the vendor’s report, the mean diameter of the QDs was 18 ( 4 nm (determined by dynamic light scattering), which is roughly consistent with our results. The surface potential (Figure 1b) image presents the contrast based on the difference in the surface potential of the QDs from that of the bare silicon substrate (grounded to 0 V). The black color of the QDs indicates that they were negatively charged in open air, which agrees with the previous literature.38 Quantitative measurements of size and surface potential were indicated by the cross-section profiles taken along the red dashed

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Figure 1. High-resolution images of the topography (a) and surface potential (b) of QDs. The cross-section profiles of the topography (c) and surface potential (d) were taken along the directions marked with the red dashed lines in (a) and (b), respectively. The white scale bars on the bottom left in (a) and (b) indicates a length of 400 nm. Histogram of the surface potential distribution of QDs (e). Size dependence of surface potential of QDs (f). The statistical distribution was obtained by randomly selecting 50 QDs with red dotted circles from Figure 1b and calculating the mean/standard deviation.

lines randomly drawn across the topography and surface potential images. The cross-section profile of the topography (Figure 1c) shows that the width of the QDs matched the abovementioned diameter, whereas their height was approximately 8 nm, less than their mean diameter. This artifact is commonly observed in tapping mode AFM, probably due to the hysteresis of the servo system as discussed by Velegol et al.39 The crosssection profile of the surface potential (Figure 1d) reveals that the QDs had a negative surface potential, which is previously reported.40 A statistical measurement of the surface potential with 50 randomly selected QDs was conducted (Figure 1e), and the mean surface potential was -154 mV with a standard deviation of 35 mV. Furthermore, the surface potential is also strongly dependent on particle diameter. QDs marked with red dotted circles in Figure 1b were selected to examine the size dependence of surface potential and the result was shown in Figure 1f. This unique electrical property may be of interest for applications of QDs and may also impact the QD particle stability in aqueous environments as well as interactions with their surroundings.14 Not only present in QDs, this sizedependent property of surface potential is found in many other metal-based nanoparticles (e.g., Fe2O3, CeO2, Au, and Ag) using KFM (results are not shown in this manuscript). This mechanisms of this size dependency remains an open question but

can potentially be interpreted by using nonlinear PoissonBoltzmann equation in the classic electrochemical knowledge in water chemistry.41,42 3.2. Characterization of Double-Stranded Homopolyers. Double-stranded DNA homopolymers [poly(dA) · poly(dT) and poly(dG) · poly(dC)] have repetitive sequences, either A · T or G · C. These simple and distinct sequences allow us to investigate the sequence dependence of DNA binding with QDs. Poly(dA) · poly(dT) was also examined by KFM prior to QD exposure, and the morphology (Figure 2a,b), surface potential (Figure 2c), and phase (Figure 2d) were simultaneously acquired. As shown in the topography images (Figure 2a,b), poly(dA) · poly(dT) has a dense network structure. The molecules varied in length from tens of nanometers to a few micrometers and had widths of approximately 10-35 nm. Some of the molecules were branched, and some aggregates had topologies that could not be followed easily. These results appear to be similar to or comparable with those reported in the literature.43 At a higher zoom, more evident morphology and surface potential (Figure 2b,c) reveals a highly tangled network with negative surface charge. The phase image (Figure 2d) measures the difference between the phase angle of the excitation and of the tip response and presents the resulting contrast due to the heterogeneity of these homopolymers, such as variations in compositional, adhesive, and

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Figure 2. High-resolution images of the topography (a,b), surface potential (c), and phase image (d) of poly(dA) · poly(dT) DNA prior to exposure to QDs. (e,f) The cross-section profiles of the topography and surface potential taken along the directions marked with the red dashed lines in (b) and (c), respectively. The white scale bars on the bottom left indicate a length of 1000 nm in (a) and 245 nm in the other images.

mechanical properties.44 Similar to the QDs, the precise measurement of size and surface potential were obtained from the cross-section profiles (Figure 2e,f) taken along the red dashed lines randomly drawn across the topography and surface potential images in Figure 2a,b. The height of poly(dA) · poly(dT) was approximately 3-5 nm whereas the width ranged from 4 to 25 nm. The surface potential of poly(dA) · poly(dT) ranged from -40 to -90 mV. This negative electrical property is comparable with previous findings,45,46 which used the PoissonBoltzmann equation to calculate the electrostatic properties of DNA in solution.46 The mechanisms of the Kelvin method are well interpreted with metal and semiconductor materials with known work functions. However, the mechanisms of applying Kelvin method on nonmetallic materials (e.g., alkanes, proteins, and DNA) are not well understood yet.47-49 Like poly(dA) · poly(dT), poly(dG) · poly(dC) had dense network structures (see Supporting Information Figure S1a), was negatively charged, and had a similar size. However, the surface potential was a bit lower and ranged from -90 to -125 mV. Another subtle difference was that poly(dG) · poly(dC) presented a pattern of surface potential on its linear structure (Supporting Information Figure S1c), that was barely visible in the topog-

raphy image (Supporting Information Figure S1b) but striking in the phase image (Supporting Information Figure S1d). This pattern may arise from the helical formation of multistranded DNA ribbons, a highly twisted structure commonly found in proteins and DNA.50 3.3. Interactions between QDs and Double Stranded Homopolyers. More interesting to study than the natural characteristics of the two homopolymers is whether or how QDs interact with them. To this end, we examined the two homopolymers with KFM after exposure to 8 nM QDs. Our results show that the homopolymer DNA strands changed their morphologies dramatically upon binding with QDs. However, we did not compare different QD exposure concentrations because KFM requires optimized concentrations of QDs for good contrast. Therefore, our findings were slightly different from the studies of An et al., who reported that exposure to high concentrations of carbon NPs caused aggregation of DNA.28 In contrast, our results in Figure 3 show that poly(dA) · poly(dT) formed pearl-like spheres with diameters of approximately 300 nm (see particularly the higher zoom in Figure 3b). The surface potential (Figure 3c) and phase (Figure 3d) images seem to exhibit sharper contrast than the topography

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Figure 3. High-resolution images of the topography (a,b), surface potential (c), phase image (d) of poly(dA) · poly(dT) DNA after exposure to QDs and (e,f) the cross-section profiles of the topography and surface potential taken along the directions marked with the dashed red lines in (b) and (c), respectively. The white scale bars on the bottom left indicate a length of 500 nm in (a) and 180 nm in the other images.

image. As measured along the red dashed line, these spheres carried a slightly lower surface potential (approximately -100 mV) than the original poly(dA) · poly(dT) (up to -90 mV). The phase image provided much better contrast for resolving the background particles (marked by the red arrow) than the topography image. The in Vitro interaction of poly(dG) · poly(dC) with QDs was similar to that of poly(dA) · poly(dT). The pearl-shaped particles were observed (Supporting Information Figure S2a); their nanoparticle-like features can be observed in contrast to Supporting Information Figure S2b, especially at higher zoom in Supporting Information Figure S2b-d. These particles likely were QDs, judging from their sizes. The cross-section profiles of topography (Supporting Information Figure S2e) and surface potential (Supporting Information Figure S2f) support this and indicate that the particles had sizes ranging from 20 to 100 nm, while the surface potential varied from -60 to -135 mV, which covers the whole range of surface potentials of QDs and DNA. In summary, the binding of QDs made two types of homopolymer DNA shift in morphology from tangled networks to pearl-like particles, and no sequence-dependent binding was observed. One may doubt that QD aggregation caused the

spherical DNA patterns. However, our previous study suggested that the carboxyl group in thioglycolic acid coating (as the capping ligands) of QDs had a pKa of 3.67,14,51 and thus, only after the pH drops to 3 or less may QDs aggregate. In contrast, at pH 5 or higher (the pH level of the DNA buffer is approximately 7), more than 95% of the capping ligands were deprotonated and the relatively high negative surface charge on the QDs made them stable in water. Even if QDs aggregated in the presence of the homopolymer DNA due to the mechanism of bridging coagulation,52 the outside surfaces of these large aggregates must have been covered by the homopolymer DNA, because the original QDs had a mean negative potential of -154 mV, whereas these large spheres had a lower magnitude in surface potentials (approximately -100 mV, see Figure 3f and Supporting Information Figure S2f). Therefore, KFM is capable of not only mapping nanoscale topography but also recognizing local entities based on surface potential information. 3.4. Interactions between QDs and the DNA Extracted from E. coli Cells. Since QDs have been reported to translocate themselves into cells via many pathways (e.g., internalization53,54 or diffusion55,56), QDs can potentially permeate cells and interact with genomes unintentionally and unpredictably. Thus, it would

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Figure 4. High-resolution images of topography (a) and surface potential (b) of DNA extracted from E. coli cells before exposure to QDs. (c,d) The cross-section profiles of the topography and surface potential taken along the directions marked with the dashed red lines in (a,b), respectively. The white scale bars on the bottom left indicate a length of 250 nm.

be interesting to investigate the in vivo binding between QDs and genomes. Here we characterized original DNA extracted from E. coli cells with and without exposure to QDs. Figure 4 shows that the original DNA ranged from 0.3 to 2 µm in length and 10-40 nm in width. A known DNA structure in which two DNA chains are tangled together is indicated by a red arrow in Figure 4a. The DNA strands were darker than the background in Figure 4b, suggesting that the DNA was negatively charged. The cross-section profiles of height and surface potential along the red dashed lines in Figure 4c,d reveal that the peak heights of these strands were approximately 1.5-3.5 nm, which corresponds to a potential of approximately -50 to -90 mV. The surface potential of E. coli DNA thus is comparable with the above measurements on homopolymer DNA [poly(dA) · poly(dT) and poly(dG) · poly(dC)]. After exposure of E. coli to QDs for 12 h, total DNA was extracted from the living cells and analyzed with KFM. Interestingly, most genomic DNA samples were not found to form pearl-like spheres like the homopolymer DNA did. Only a small portion of the DNA formed particle-shaped clusters, as indicated by red arrows in the topography image of a typical DNA sample (Figure 5a). In Figure 5b, a linear DNA strand seemed to bind with QDs; to confirm this, a red dotted line was drawn across their apexes to reveal their height and surface potential profiles. In the cross-section profiles (Figure 5c,d), gray and red bars mark the normal ranges of height and surface potential for QDs and DNA. The large particle bound to the linear DNA had the highest height peak (#1) in the typical QD range (see Figure 1), whereas the lower peak (#2) next to the #1 peak had a height in the typical DNA range (see Figures 2 and 4). The #1 peak might be one large QD (or an aggregate of smaller QDs) or QDs conjugated with DNA. The #2 peak should represent linear DNA. The surface potentials in Figure 5d further support the above speculation and the #1 peak had a potential of approximately -200 mV, which is within the typical surface potential range of a similarly sized QD. The #2 peak had a

potential of approximately -100 mV, which is in the typical DNA surface potential range. The deviation between the in vivo binding of DNA with QDs and the in vitro experiments with homopolymer DNA may be caused by the partitioning of QDs onto other biomolecules such as proteins in the cytoplasm. In addition, during the DNA extraction and purification, weakly bound QDs might be removed from the DNA molecules and the deformed DNA changed back to its original structure. Therefore, the in vivo and in vitro DNA binding were different. Monitoring of the in situ binding of QDs with cellular DNA may be accomplished with other techniques such as confocal microscopy but its magnification and resolution are inferior to KFM or AFM. 3.5. Possible Interaction Mechanisms between QDs and DNA. On the basis of observations from the KFM study, QDs could bind with DNA in Vitro and in ViVo. The possible scenarios of QDs-DNA interaction mechanisms are outlined in Figure 6 and these potential mechanisms are highly related to the potential genotoxicity of QDs. Double-stranded DNA is composed of sugar-phosphate backbones and nitrogenous bases (adenin, guanine, thiamin, and cytosine). As measured by KFM, DNA molecules were negatively charged clusters; due to the electrostatic interactions, QD-DNA attachments thermodynamically unfavorable as QDs are also negatively charged. However, this attachment did occur due to other possible specific interactions.57 For example, QDs may bind to DNA through hydrogen binding and the energy released may promote the surface construction of DNA.28 As illustrated in Figure 6, the carboxylic groups (-COO-) of QDs form hydrogen bonds with the phosphate groups (PO4-) of DNA. This hydrogen bonding leads to the first and second interaction mechanisms in Figure 6. The linear attachment of QDs to DNA in the first mechanism is supported by Figure 5. The second mechanism (QDs are surrounded by DNA molecules) is just our observed consequence in the in vitro binding experiments with homopolymer DNA.

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Figure 5. High-resolution images of topography (a) and surface potential (b) of DNA extracted from E. coli cells after the exposure to QDs. Cross-section profiles of the topography and surface potential were taken along the directions marked with the dashed red lines in (a) and (b), respectively. The white scale bars on the bottom left indicate a length of 300 nm. The gray bar on the Y-axis in (c) indicates the height range of QDs obtained from Figure 1, whereas the red bar indicates the DNA height range obtained from Figures 2 and 4. The gray bar on the Y-axis in (d) indicates the surface potential range of QDs obtained from Figure 1, whereas the red bar indicates the DNA surface potential range obtained from Figures 2 and 4.

confirm the third and fourth mechanisms experimentally. These binding mechanisms apparently cause DNA to cross-link and form more complicated levels of organization, which may interfere with or prevent the DNA from extending during the replication process. This disruption in DNA replication disturbs the gene expression and normal functioning of cells. Thus, the exposure of QDs could cause adverse biological effect.11,16,40 Our study with E. coli cells exposed to QDs also supports this and Supporting Information Figure S3 shows that as the exposure concentration of QDs increased, cellular growth was inhibited, as indicated by the lower values of optical density (OD600), a measure of biomass concentration based on adsorbance of the cell culture at the wavelength of 600 nm. Although cellular inhibition may have multiple causes (i.e., oxidative stress) besides DNA binding with QDs, the two could be correlated because as mentioned above the disruption of DNA replication can lead to cellular mutation or death.59 4. Conclusions

Figure 6. A representative DNA vector interaction with QDs and four possible interaction mechanisms of their interactions (relative sizes are not to scale).

Similar to the second mechanism, in the third mechanism QDs also likely bended the linear DNA owing to the highly twisted structure and bending property of DNA.58 The fourth potential interaction is the embedding of QDs into the nitrogenous bases of DNA. However, we are currently unable to

We have demonstrated the outstanding performance of KFM in mapping the topography, morphology, and surface potential of QDs, homopolymer DNA, and E. coli DNA. Both in vitro and in vivo binding between DNA and QDs was visualized successfully with KFM for the first time, and homopolymer DNA transformed into pearl-like spheres after exposure to QDs. Such morphological changes in DNA were observed for the first time with KFM and these structural conformations could change the nanoelectrical properties of DNA and cause the growth inhibition as we observed in the E. coli cells. KFM was shown to be a powerful technique for imaging, quantifying, and recognizing interacting entities at the nanoscale. Important advantages include the high-resolution imaging and identification of NPs in biological matrices and less labor for sample

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preparation (e.g., no labeling is required) compared to confocal microscopy. KFM potentially can be used in surface characterization of nanomaterials and in resolution of nanobio interfacial phenomena such as type-specific binding between NPs and biomolecules (DNA or proteins). The imaging and quantification capabilities of KFM provide a unique angle for both applications and implications of nanotechnology research at the molecular level. Acknowledgment. This study was partially supported by the U.S. Environmental Protection Agency Science to Achieve Results Program Grants (RD-83385601 and RD-831713) and Semiconductor Research Corporation (SRC)/ESH grant. Supporting Information Available: Additional figures. This material is available free of charge via the Internet at http:// pubs.acs.org. References and Notes (1) Alivisatos, A. P. Science 1996, 271, 933. (2) Murray, C. B.; Kagan, C. R.; Bawendi, M. G. Annu. ReV. Mater. Sci. 2000, 30, 545. (3) Klabunde, K. J. Nanoscale materials in chemistry; John Wiley & Sons, Inc.: New York, 2001. (4) Chen, L.-D.; Liu, J.; Yu, X.-F.; He, M.; Pei, X.-F.; Tang, Z.-Y.; Wang, Q.-Q.; Pang, D.-W.; Li, Y. Biomaterials 2008, 29, 4170. (5) Walther, C.; Meyer, K.; Rennert, R.; Neundorf, I. Bioconjugate Chem. 2008, 19, 2346. (6) Zhang, C. Y.; Hu, J. Anal. Chem. 2010, 82, 1921. (7) Zhang, C. Y.; Yeh, H. C.; Kuroki, M. T.; Wang, T. H. Nat. Mater. 2005, 4, 826. (8) Luque, A.; Martı´, A.; Nozik, A. J. MRS Bulletin 2007, 32, 236. (9) Ron, H. EnViron. Health Perspect. 2006, 114, 165. (10) Priester, J. H.; Stoimenov, P. K.; Mielke, R. E.; Webb, S. M.; Ehrhardt, C.; Zhang, J. P.; Stucky, G. D.; Holden, P. A. EnViron. Sci. Technol. 2009, 43, 2589. (11) Choi, A.; Brown, S.; Szyf, M.; Maysinger, D. J. Mol. Med. 2008, 86, 291. (12) Cho, S. J.; Maysinger, D.; Jain, M.; Ro¨der, B.; Hackbarth, S.; Winnik, F. M. Langmuir 2007, 23, 1974. (13) Chang, E.; Thekkek, N.; Yu, W. W.; Colvin, V. L.; Drezek, R. Small 2006, 2, 1412. (14) Zhang, Y.; Chen, Y.; Westerhoff, P.; Crittenden, J. C. EnViron. Sci. Technol. 2007, 42, 321. (15) Koeneman, B. A.; Zhang, Y.; Hristovski, K.; Westerhoff, P.; Chen, Y.; Crittenden, J. C.; Capco, D. G. Toxicol. in Vitro 2009, 23, 955. (16) Schneider, R.; Wolpert, C.; Guilloteau, H.; Balan, L.; Lambert, J.; Merlin, C. Nanotechnology 2009, 20, 225101. (17) Verma, A.; Stellacci, F. Small 2010, 6, 12. (18) Qi, L.; Gao, X. Expert Opin. Drug DeliVery 2008, 5, 263. (19) Mitchell, G. P.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1999, 121, 8122. (20) Medintz, I. L.; Uyeda, H. T.; Goldman, E. R.; Mattoussi, H. Nat. Mater. 2005, 4, 435. (21) Parak, W. J.; Pellegrino, T.; Micheel, C. M.; Gerion, D.; Williams, S. C.; Alivisatos, A. P. Nano Lett. 2003, 3, 33. (22) Kimura-Suda, H.; Petrovykh, D. Y.; Tarlov, M. J.; Whitman, L. J. J. Am. Chem. Soc. 2003, 125, 9014. (23) Yan, H.; Park, S. H.; Finkelstein, G.; Reif, J. H.; LaBean, T. H. Science 2003, 301, 1882. (24) Nel, A. E.; Madler, L.; Velegol, D.; Xia, T.; Hoek, E. M. V.; Somasundaran, P.; Klaessig, F.; Castranova, V.; Thompson, M. Nat. Mater. 2009, 8, 543.

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