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Photoluminescent Carbon Dots as Biocompatible Nanoprobes for Targeting Cancer Cells in Vitro Qin Li,*,† Tymish Y. Ohulchanskyy,‡ Ruili Liu,*,§ Kaloian Koynov,| Dongqing Wu,| Andreas Best,| Rajiv Kumar,‡ Adela Bonoiu,‡ and Paras N. Prasad*,‡ Department of Chemical Engineering, Curtin UniVersity of Technology, Perth, Western Australia 6845, Australia, Institute of Laser, Photonics and Biophotonics, UniVersity at Buffalo, The State UniVersity of New York, Buffalo, New York 14260, School of EnVironmental and Chemical Engineering, Shanghai UniVersity, Shangda Road 99, Shanghai 200444, People’s Republic of China, and Max Planck Institute for Polymer Research, Mainz 55128, Germany ReceiVed: December 4, 2009; ReVised Manuscript ReceiVed: April 27, 2010
Carbon nanoparticles become photoluminescent upon surface passivation with oligomeric polymer chains. In this work, the dependence of the carbon dots photoluminescent properties on the passivation polymer selection has been demonstrated by conjugating polyethylene glycol (PEG) chains, polyethylenimide-co-polyethylene glycol-co-polyethylenimide copolymer, and 4-armed PEG molecules, respectively. The cytotoxicity and cellular internalization of the resulting three types of photoluminescent nanoformulations of carbon dots, named CD2, CD3, and CD4, were evaluated. These nanoformulations exhibited no apparent cytotoxicity on their own and were shown to successfully target cancer cells by conjugation with transferrin. The implication to the use of carbon dots as biocompatible optical nanoprobes for in vitro cancer diagnostics is discussed. Introduction The importance of fluorescent nanoprobes in biomedical research and practice has been rapidly increasing as a converged outcome of fast developments in fluorescence microscopy, laser technologies, and nanotechnology.1 There has always been a constant search for benign “nanolanterns” that can be used for studying intracellular transport and biochemical interactions, which ultimately can be used for disease detection and therapy.2 It was first discovered by Sun et al.3 that laser-ablated, amorphous carbon nanoparticles can emit in the visible spectral range upon surface functionalization with polymer chains. These carbon dots (CDs) as illustrated in Scheme 1 have demonstrated significant potential as a new class of photoluminescent nanoparticles. Compared to the semiconductor-based quantum dots, which have been widely explored particularly in biological applications,4–7 CDs have the advantage in their chemical inertness, versatile surface chemistry, and potentially low cytotoxicity.3,8 A unique property of this class of multicolored nanoparticles is that the bare carbon nanoparticles are not luminescent; however, upon covalent coupling with polymer chains, the carbon/polymer composites become photoluminescent.3,8,9 Though the exact mechanism of photoluminescence (PL) remains elusive, it has been hypothesized that it is the surface energy traped on the bare CD surface, which becomes emissive after passivation.3 This feature is similar to what has been observed on the photoluminescence of carbon nanotubes,10 where the emission spectra can be altered by the variation of functional moieties and density.11 However, carbon nanotubes may induce undesired effects to the biological entities, which have been attributed to the metal catalyst residue or the “asbestos
effect”.12 Moreover, the superb transmembrane ability of the carbon nanotubes13 may limit its specificity for targeted delivery. Carbon nanodiamonds14,15 are also known to be photoluminescent. Nitrogen-doped nanodiamonds have recently attracted considerable attention owing to their superior quantum yield and the ability to be internalized by cells.16,17 However, their size (>10 nm) so far synthesized may become problematic for clinical applications, because they are too large to be cleared from the body by renal excretion,18 and they are not degradable. Recently, we reported a novel aqueous route to synthesizing nanosized (1.5-2.5 nm) CDs by employing surfactant-modified silica spheres as carriers.8 Similar to the CDs prepared by laser ablation,3,19 they become photoluminescent upon conjugating polymer chains, with a quantum yield up to 15%, and have excellent biocompatibility.8,19 The objective of this work is 2-fold: (i) to demonstrate that the photoluminescent properties of the CDs can be conveniently tuned by using nanoformulations with varying functional polymer chains and (ii) to investigate the potential of carbon dots as biocompatible nanoprobes for targeting cancer cells in Vitro. SCHEME 1: Schematic of a Carbon Dot Structure
* To whom correspondence should be addressed. E-mail: Q.Li@ curtin.edu.au,
[email protected], and
[email protected]. † Curtin University of Technology. ‡ University at Buffalo. § Shanghai University. | Max Planck Institute for Polymer Research.
10.1021/jp911539r 2010 American Chemical Society Published on Web 06/24/2010
Photoluminescent Carbon Dots Experimental Methods Materials. Tetraethyl orthosilicate (TEOS), phenol, formalin solution (37 wt %), NaOH, HNO3, Na2CO3, and poly(propylene oxide)-co-poly(ethylene oxide)-co-poly(propylene oxide) triblock copolymer Pluronic F127 (MW ) 12 600, EO106PO70EO106) were purchased from Fluka. Diamine-terminated oligomeric polymeric poly(ethylene glycol), H2NCH2(CH2CH2O)nCH2CH2CH2NH2 (average n ≈ 35, PEG1500N) and amine-terminated, 4-armed poly(ethylene oxide) (PEO) were purchased from Sigma-Aldrich. Triblock copolymer consisting of poly(ethylenimide)-co-poly(ethylene glycol)-co-poly(ethylenimide) (PEI-PEG-PEI, MW ) 5k-5k-5k) was kindly provided by A. Taubert et al.20 Transferrin and 1-ethyl-3-[3dimethylaminopropyl]carbodiimide hydrochloride (EDC) were purchased from Sigma Aldrich. All chemicals were used as received without any further purification. The cancer cell lines HeLa cells and Pancrea-1cells were obtained from ATCC. The Cell Proliferation Assay (Pomega GPR G5421) and MTS cytotoxicity assay were both acquired from Promega. Preparation of Different Types of Photoluminescent CDs. The bare CDs were prepared as described previously, using the aqueous route with silica spheres as carriers.8 After being treated with concentrated nitric acid for introducing carboxyl groups, the nascent CDs were mixed with three types of polymers, respectively: polymer 1, poly(ethylene glycol), H2NCH2(CH2CH2O)nCH2CH2CH2NH2 (average n ≈ 35, PEG1500N); polymer 2, a triblock copolymer consisting of poly(ethylenimide)-b-poly(ethylene glycol)-b-poly(ethylenimide) (PEI-PEGPEI, 5k-5k-5k); and polymer 3, a 4-arm, amine-terminated PEG (4-arm PEG, MW ) 10k). Each mixture suspension of bare CDs and excessive polymer was heated at 120 °C under reflux for 72 h for surface passivation. Subsequently, the optically transparent and photoluminescent CD suspensions were purified by dialyzing against Milli-Q water with a cellulose ester membrane bag. For CD2, a MWCO 3500 membrane bag was used; while for CD4, a MWCO 14 000 dialysis membrane was applied. It should be noted that CD3 was not purified with the dialysis method because of the difficulty in separation due to the large size of the PEI-PEG-PEI molecules. The final three nanoformulations with the three different polymer coatings were named as CD2 (CD-PEG1500N), CD3 (CD-PEI-PEG-PEI), and CD4 (CD-4 arm PEG). Optical Characterizations. Optical absorption and PL spectroscopy were used to examine the spectral properties of the CDs. UV-vis absorption spectra were recorded with a Shimadzu 3600 UV-vis-NIR spectrophotometer, using a quartz cuvette with a 1 cm path length. PL excitation and emission spectra were acquired on a Fluorolog-3 spectrofluorimeter (Jobin Yvon, Longjumeau, France). Dynamic light scattering (Brookhaven 90PLUS with ZetaPALS option) was used to determine the ζ-potential of the three nanoformulations of carbon dots. Fluorescence Correlation Spectroscopy. The concentration and hydrodynamic diameter of the photoluminescent particles in the suspensions were characterized by fluorescence correlation spectroscopy (FCS). FCS was performed with a commercial setup (Carl Zeiss, Germany), consisting of the module ConfoCor 2 and an inverted microscope model Axiovert 200.21,22 The QDs were excited by an argon laser (λ ) 488 nm) and the emission was collected after filtering with a LP505 long pass filter. The eight-well, polystyrene chambered cover-glass (Lab-Tek, Nalge Nunc International) was used as a sample cell. For each solution, a series of ten measurements with a total duration of 5 min were performed. From the measured temporal fluctuations of
J. Phys. Chem. C, Vol. 114, No. 28, 2010 12063 the fluorescence intensity, δI(t), an autocorrelation function G(τ) ) 1 + 〈δI(t)δI(t + τ)〉/〈I(t)〉2 was evaluated. As has been shown theoretically for an ensemble of identical freely diffusing fluorescence species, G(τ) has the following analytical form21,23
G(τ) ) 1 +
(
1 τ 1+ N τD
)( -1
1+
τ S τD 2
)
-1/2
(1)
where N is the average number of fluorescent molecules/particles in the observation volume V that is directly related to their concentration, τD is the lateral diffusion time that a particle stays in the volume, and S ) z0/r0 is the ratio of axial to radial dimensions of V (S ≈ 6 in our experiment). Furthermore, the diffusion time, τD, is related to the respective diffusion coefficient, D, through21,23
D ) r02 /4τD
(2)
The experimentally obtained G(τ) values were fitted with eq 1 yielding the diffusion time τD, and subsequently the diffusion coefficient of the fluorescent species (eq 2). Finally, the average particle size was determined by the Stokes-Einstein relation.22 As the value of r0 depends strongly on the specific characteristics of the optical setup and the refractive index of the studied samples, a calibration was performed by measurement of the characteristic diffusion time of a reference fluorescent tracer with known diffusion coefficient, i.e., Rhodamine 6G. Conjugation of CDs with Transferrin. For targeted in Vitro imaging studies, the NH2 terminated CDs were conjugated with transferrin by using EDC chemistry. In a typical protocol, 0.5 mg of transferrin was added to 0.5 mL of ultrapure water followed by the addition of 25 µL of 0.1 M EDC solution and the mixture was gently stirred for 30 min. Next, 1 mL of the nanoparticles was added and the mixture was incubated at room temperature for 2 h to allow the protein to covalently bond to the NH2 groups of the CDs. Studies of Cellular Uptake and Targeting. For in Vitro imaging, the HeLa cell line was cultured in Dulbecco minimum essential media (DMEM) with 10% fetal bovine serum (FBS), 1% penicillin, and 1% amphotericin B. The day before nanoparticles treatment, cells were seeded in 35 mm culture dishes. On the treatment day, the cells, at a confluency of 50-60%, in serum-supplemented medium were treated with the CD nanoparticles at a specific concentration (100 µL per 1 mL of medium) for 2 h at 37 °C. Cellular Imaging. Confocal microscopy images were obtained with a Leica TCS SP2 AOBS spectral confocal microscope (Leica Microsystems, Wetzler, Germany) using laser excitation at 442 nm. All confocal images, which were compared, were obtained at the same parameters of laser power, confocal pinhole, gain, offset, and scanning speed. Cell Viability Assay. The cellular cytotoxicity of CD2, CD3, and CD4 was tested on Hela cells. The cell viability assay was performed with a CellTiter 96AQueous Non-Radioactive Cell Proliferation Assay (Pomega GPR G5421) based on reduction of a tetrazolium component [3-(4,5-dimethylthiazol-2-yl)-5-(3carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt; MTS] into an insoluble formazan product by the mitochondria of viable cells.24 HeLa cells were seeded in a 96 well plate (∼10 000 cells/mL/well) and maintained in culture medium for 24 h at 37 °C. Then 10 µL of CD2/CD3/CD4 was added to every well containing 100 µL of medium and cells were
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Figure 1. Absorption spectra of CD2, CD3, and CD4 (a) and PL emission spectra of CD2 (b), CD3 (c), and CD4 (d). The inserted photos were taken when the suspensions were excited with an UV lamp (365 nm).
Figure 2. PL excitation spectra of CD2 (a), CD3 (b), and CD4 (c). PL was monitored at the different wavelengths shown in the insets.
Photoluminescent Carbon Dots
Figure 3. Cell viability (MTS assay) of HeLa cells following treatment with CD2, CD3, and CD4 for 24 h. The results are the mean ( SD of three separate experiments.
incubated for another 24 h. After this, the cells were carefully rinsed, followed by treatment with MTS reagent for 2 h. The produced formazan was quantified by measurement of its absorbance at 490 nm, using an Elisa plate reader. Results and Discussion The particle morphology was observed by TEM and AFM (see Supporting Information), showing particle size in the range of 1.5-3 nm. The concentrations and the average hydrodynamic diameters of the as-prepared CDs were obtained by FCS as 1.1 µM, 2.6 nm for CD2; 1.4 µM, 1.84 nm for CD3; and 250 nM, 3.6 nm for CD4. Apparently, the mixture of polymer 3 and nascent CDs yielded a 5-fold lower amidation efficiency, possibly due to steric hindrance caused by the large size of polymer 3, the 4-armed 10 kD PEG. It is particularly curious
J. Phys. Chem. C, Vol. 114, No. 28, 2010 12065 with the hydrodynamic size of CD3, being the smallest despite the conjugation with a MW 15k copolymer. Here we suppose that the copolymer-CD composite of CD3 may be of a certain irregular shape, whose diffusion rate significantly deviates from the Einstein-Stokes diffusion equation that is only applicable to spheres. Therefore, the hydrodynamic size of CD3 here is invalid. Optical Properties. All three nanoformulations of CDs showed broad absorption in the UV-vis range. As seen in Figure 1a, CD3 suspension absorbs much stronger than CD2 and CD4. There is also a shoulder on the CD3 absorption curve at about 370 nm, which is different from the monotonically falling curves for CD2 and CD4. The difference in absorbance is much higher than would be expected from the difference in the CD concentration, which leads us to assume that the passivation of the CDs surface with PEI-PEG-PEI results in the formation of a higher amount of the absorbing centers. Under excitation at 365 nm, both CD2 and CD3 suspensions exhibited blue luminescence, while emission from CD4 appears yellowish (see photos in Figure 1). The PL spectra of CD2 and CD3 are shown in Figure 1, panels b and c, respectively. They both are multicolored as reported previously,3,8 where the emission maxima shifted bathochromically as the excitation wavelength increased stepwise from 320 to 600 nm. There are also noticeable differences between the PL of CD2 and CD3. Whereas CD3 seems to emit more intensively than CD2, one should note that the absorbance of CD3 is much higher. Furthermore, the most intensive PL from CD3 appears under 440 nm excitation and has a maximum at ∼510 nm, while CD2 photoluminescence is the most intensive under excitation at 360 nm, with the peak positioned at 460 nm (Figure 1). It is worth
Figure 4. Carbon dots (a) CD2, (b) CD3, (c) CD4, (d) Tf-CD2, (e) Tf-CD3, and (f) Tf-CD4 with internalization after 2 h of incubation with HeLa cells. Photoluminescence (shown as green) and transmission images are merged.
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Figure 5. Targeting HeLa cells with transferrin conjugated carbon dots. Comparison of cells treated with (a) CD3 (top transmission and PL images) versus CD3-Tf (middle transmission and PL images) versus CD3-Tf incubated with cells pretreated with free Tf for 1 h (bottom transmission and PL images) and cells nonpretreated with free Tf; (b) CD4-Tf incubated with cells pretreated with free Tf for 1 h (top transmission and PL images) and cells nonpretreated with free Tf (bottom transmission and PL images).
noting that the PL emission spectra of CD4 (Figure 1d) appear very different from those of CD2 and CD3. The lowest light absorption and PL intensities are partly due to the lower concentration of CD4. Nevertheless, the most interesting observation was that when CD4 was excited at 320 nm, the PL spectrum was extremely broad, peaking at around 550 nm, with the highest intensity among the series PL spectra. Second, when increasing the excitation wavelength, both the width and the intensity of the PL emission spectra decreased, and the maxima position is even blue-shifted (up to ∼510 nm) when the excitation wavelength is increased from 320 to 400 nm. When the excitation is further tuned toward longer wavelengths (>400 nm), the PL emission band starts to move bathochromically up to ∼650 nm. This overall complex behavior of the PL emission spectra is apparently associated with a variety of emitting centers present in the CD suspensions. In support of this, the decays of PL emission are nonmonoexponential for all three types of CD suspensions, with the average lifetime in the range of a few nanoseconds (see the Supporting Information). An inspection of the PL excitation spectra (Figure 2) confirms the existence of multiple types of emitting centers; however, they can be assigned to a few distinct types. The PL excitation band of CD3 appears as continuously shifting but also can be presented as a combination of two PL excitation bands with maxima at ∼410 and ∼460 nm. The PL excitation spectra of CD2 and CD4 are
different from those of CD3, but similar to each other. In particular, one can distinguish the PL excitation bands which are common to both CD2 and CD4, with maxima at ∼370 nm, two other peaks at ∼420-430 and ∼500 nm. It is important to note that all the differences in the optical properties of CD2, CD3, and CD4 are merely caused by the difference in polymer chains bound to the nanoparticle surface, despite that all three types of polymers were coupled on the nascent CDs by an identical amidation procedure. Similar functionalization dependence has been observed on the photoluminescence of carbon nanotubes (CNT), which was attributed to the improved CNT dispersity and surface passivation quality.11 The PL excitation and emission spectra of the CD suspensions are determined by the type of polymer conjugated to CDs. Thus, a presence of PEI leads to the difference between the PL excitation and emission spectra of CD3 and the other two types of CDs, which do not contain PEI. Similarly, a structural difference between PEG and 4-arm PEG in CD2 and CD4, correspondingly, determines the formation of the differing nanoparticle-polymer conjugation sites, which are specific to the type of polymer and responsible for optical absorption and photoluminescence. The FCS measurement also reveals the single nanoparticles brightness expressed in photon counts per particle for a given excitation laser intensity. CD4 has shown 20 kHz/particle compared to 4.4 of CD2. The causes of CD4’s
Photoluminescent Carbon Dots highest single particle photon count as well as having the largest Stokes shift when excited by UV light remain elusive and are currently under further investigation. Cytotoxicity. The CDs were introduced into the HeLa cell cultures to evaluate their potential as bioimaging agents. The cytotoxicity of the CDs was also evaluated. As can be seen in Figure 3, CD2, CD3, and CD4 demonstrate low cytotoxicity. It is worthwhile to point out that the concentration used in this cytotoxicity experiment is twice as high as the CD concentration used for cell imaging. Targeting Cancer Cells. For applications of both disease detection and drug delivery, it is desirable that the photoluminescent nanoparticles can be tailored to actively target a specific type of cell or a particular intracellular compartment as rapid passive cellular uptake of these nanoparticles may not necessarily be advantageous in many biomedical applications. To test the usability of the CDs nanoformulation as targeting photoluminescent nanoprobes, all three CDs were coupled with human transferrin (Tf) through carbodiimide chemistry following established protocol.25 Tf, a serum glycoprotein (80 kDa), has been known for its ability in targeting cancer cells due to overexpression of Tf receptors on cancer cell membranes.26 The PL emission spectra of the Tf conjugated CDs remain the same as before conjugation (data not shown). A 200 µL sample of CD2, CD3, CD4, Tf-CD2, Tf-CD3, and Tf-CD4 (Tf conjugated CDs are of equivalent concentrations to their corresponding parent suspensions) was added to dishes with HeLa cells, containing 2 mL of the medium. After 2 h of incubation, HeLa cells were shown to internalize the protein conjugated CD2 and CD4 much more efficiently than non-Tf-conjugated CD2 and CD4 (Figure 4a,e and 4b,f). CD3 demonstrated higher passive cellular uptake in comparison with the other two types of nonTf-conjugated CDs by the HeLa cells (Figure 4c). This outcome is consistent with the nanoparticles physical and chemical properties. Due to the excessive amine groups on the CD3 surface carried by the PEI-PEG-PEI polymer chains, CD3 is positively charged (ζ potential +3.35 mV). Therefore, CD3 is more capable of binding with the cell membrane through electrostatic interactions.25 On the other hand, although CD2 and CD4 also have amine groups on the surface due to the amine terminated PEG chains, their overall surface charges are negative; the values of ζ potential are -14.51 mV for CD2 and -2.98 mV for CD4. We attribute this negative charge to the density of polymer coverage on the surface as well as the charge negativity of the PEG segment, a widely accepted antibiofouling polymer. These cellular uptake experiments have shown that the design of surface functional groups of all three types of CDs permit efficient conjugation ability with biomolecules through carbodiimine chemistry. A selection of functional polymer groups for conjugation to the nanoparticle surface to a great extent determines the efficiency of the cellular uptake of the nanoparticles. It is worth noting that the difference in the PL signal between the cells treated with Tf-conjugated and non-Tf-conjugated CDs looks more pronounced for CD2 and CD4 than in the case of CD3. However, one can still discriminate the difference between non-Tf-conjugated and Tf-conjugated CD3, if imaging conditions (e.g., gain of the confocal PL detector) are chosen in such a way that no PL signal is visible for passive cellular uptake (non-Tf-conjugated CD3). Consequently, the preferential PL signal from the cells targeted with Tf-conjugated CD3 (active cellular uptake) would be easily seen (Figure 5b). To check whether the increased internalization of the Tfconjugated CDs was indeed mediated by the Tf receptors which
J. Phys. Chem. C, Vol. 114, No. 28, 2010 12067 were overexpressed on cancer cell membranes,26 we pretreated cells with free Tf before treating them with Tf-conjugated CDs. As seen in Figure 5, pretreatment of cells with free Tf causes a sharp decrease in the targeting efficiency for Tf-conjugated CDs, suggesting saturation of the Tf receptors with free Tf.27 These data strongly support a Tf-receptor-mediated cellular uptake of the Tf-conjugated CDs, confirming the cell targeting capability of the bioconjugated carbon dots. Conclusions In conclusion, our study demonstrates that CDs can be used as wavelength-tunable optical nanoprobes with flexibility to surface functionalization and bioconjugation. Therefore, it can readily acquire targeted delivery function when conjugated with targeting motifs, for addressing disease cells such as cancer cells. The extraordinary emission attained by conjugation of polymer chains affords an infinitive amount of tuning possibilities for improving and tailor-designing the optical properties of this new class of materials. Acknowledgment. The authors are grateful to the generosity of A. Taubert’s group at the University of Potsdam for providing the PEI-PEG-PEI block copolymer. Q.L. thanks the support of the Australian Research Council (DP0558727) and Curtin University Strategic Research Funding. P.N.P. thanks the John R. Oishei Foundation for support. R.L. gratefully acknowledges the National Natural Science Foundation of China (20903066) for financial support. Supporting Information Available: TEM and AFM characterization of the carbon dots and PL emission decays of CD2, CD3, and CD4 and the determined lifetime. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Hurtley, S. M.; Helmuth, L. Science 2003, 300, 75. (2) Michalet, X.; Pinaud, F. F.; Bentolila, L. A.; Tsay, J. M.; Doose, S.; Li, J. J.; Sundaresan, G.; Wu, A. M.; Gambhir, S. S.; Weiss, S. Science 2005, 307, 538. (3) Sun, Y. P.; Zhou, B.; Lin, Y.; Wang, W.; Fernando, K. A. S.; Pathak, P.; Meziani, M. J.; Harruff, B. A.; Wang, X.; Wang, H. F.; Luo, P. J. G.; Yang, H.; Kose, M. E.; Chen, B. L.; Veca, L. M.; Xie, S. Y. J. Am. Chem. Soc. 2006, 128, 7756. (4) Gao, X.; Cui, Y.; Levenson, R. M.; Chung, L. W. K.; Nie, S. Nat. Biotechnol. 2004, 22, 969. (5) Alivisatos, A. P. Nat. Biotechnol. 2004, 22, 47. (6) Law, W.-C.; Yong, K.-T.; Roy, I.; Ding, H.; Hu, R.; Zhao, W.; Prasad, P. N. Small 2009, 5, 1302. (7) Derfus, A. M.; Chan, W. C. W.; Bhatia, S. N. Nano Lett. 2004, 4, 11. (8) Liu, R.; Wu, D.; Liu, S.; Koynov, K.; Knoll, W.; Li, Q. Angew. Chem., Int. Ed. 2009, 48, 4598. (9) Cao, L.; Wang, X.; Meziani, M. J.; Lu, F. S.; Wang, H. F.; Luo, P. J. G.; Lin, Y.; Harruff, B. A.; Veca, L. M.; Murray, D.; Xie, S. Y.; Sun, Y. P. J. Am. Chem. Soc. 2007, 129, 11318. (10) Riggs, J. E.; Guo, Z.; Carroll, D. L.; Sun, Y.-P. J. Am. Chem. Soc. 2000, 122, 5879. (11) Lin, Y.; Zhou, B.; Martin, R. B.; Henbest, K. B.; Harruff, B. A.; Riggs, J. E.; Guo, Z. X.; Allard, L. F.; Sun, Y. P. J. Phys. Chem. B 2005, 109, 14779. (12) Lewinski, N.; Colvin, V.; Drezek, R. Small 2008, 4, 26. (13) Kostarelos, K.; Lacerda, L.; Pastorin, G.; Wu, W.; Wieckowski, S.; Luangsivilay, J.; Godefroy, S.; Pantarotto, D.; Briand, J. P.; Muller, S.; Prato, M.; Bianco, A. Nat. Nanotechnol. 2007, 2, 108. (14) Gruber, A.; Drabenstedt, A.; Tietz, C.; Fleury, L.; Wrachtrup, J.; von Borczyskowski, C. Science 1997, 276, 2012. (15) Yu, S. J.; Kang, M. W.; Chang, H. C.; Chen, K. M.; Yu, Y. C. J. Am. Chem. Soc. 2005, 127, 17604. (16) Faklaris, O.; Garrot, D.; Joshi, V.; Druon, F.; Boudou, J. P.; Sauvage, T.; Georges, P.; Curmi, P. A.; Treussart, F. Small 2008, 4, 2236. (17) Krueger, A. AdV. Mater. 2008, 20, 2445.
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(18) Choi, H. S.; Liu, W.; Misra, P.; Tanaka, E.; Zimmer, J. P.; Ipe, B. I.; Bawendi, M. G.; Frangioni, J. V. Nat. Biotechnol. 2007, 25, 1165. (19) Yang, S. T.; Wang, X.; Wang, H. F.; Lu, F. S.; Luo, P. J. G.; Cao, L.; Meziani, M. J.; Liu, J. H.; Chen, M.; Huang, Y.; Sun, Y. P. J. Phys, Chem. C 2009, 113, 18110. (20) Grumelard, J.; Taubert, A.; Meier, W. Chemical Communications 2004, 1462. (21) Rigler, R.; Elson, E. Fluorescence Correlation Spectroscopy: Theory and Applications; Springer: Berlin, Germany, 2001. (22) Koynov, K.; Mihov, G.; Mondeshki, M.; Moon, C.; Spiess, H. W.; Muellen, K.; Butt, H.-J.; Floudas, G. Biomacromolecules 2007, 8, 1745.
Li et al. (23) Haustein, E.; Schwille, P. Curr. Opin. Struct. Biol. 2004, 14, 531. (24) Cory, A. H.; Owen, T. C.; Barltrop, J. A.; Cory, J. G. Cancer Commun. 1991, 3, 207. (25) Kumar, R.; Roy, I.; Hulchanskyy, T. Y.; Goswami, L. N.; Bonoiu, A. C.; Bergey, E. J.; Tramposch, K. M.; Maitra, A.; Prasad, P. N. ACS Nano 2008, 2, 449. (26) Qian, Z. M.; Li, H. Y.; Sun, H. Z.; Ho, K. Pharmacol. ReV. 2002, 54, 561. (27) Racoosin, E. L.; Swanson, J. A. J. Cell Sci. 1992, 102, 867.
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