Reversible Self-Assembly of Nanoprobes in Live Cells for Dynamic

Subscriber access provided by TULANE UNIVERSITY

Article

Reversible Self-Assembly of Nanoprobes in Live Cells for Dynamic Intracellular pH Imaging Dingbin Liu, Bo Dong, Shuangli Du, Chunxiao Wang, Haohao Fu, Qiang Li, Nannan Xiao, Jie Yang, Xue Xue, and Wensheng Cai ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.8b07054 • Publication Date (Web): 07 Feb 2019 Downloaded from http://pubs.acs.org on February 7, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Reversible Self-Assembly of Nanoprobes in Live Cells for Dynamic Intracellular pH Imaging Bo Dong,1 Shuangli Du,1 Chunxiao Wang,2 Haohao Fu,1 Qiang Li,1 Nannan Xiao,2 Jie Yang,1 Xue Xue,2 Wensheng Cai,1 and Dingbin Liu*,1,2 1

College of Chemistry, Research Center for Analytical Sciences, and Tianjin Key Laboratory of

Molecular Recognition and Biosensing, Nankai University, Tianjin 300071 (China) 2 State

Key Laboratory of Medicinal Chemical Biology, Nankai University, Tianjin 300071 (China)

*Correspondence: [email protected] ABSTRACT: Self-assembly is a powerful tool to organize the elementary molecular units into functional nanostructures, which provide reversible stimulus-responsive systems for a variety of purposes. However, the ability to modulate the reversible self-assembly in live systems remains a great challenge owing to the chemical complexity of intracellular environments which often damage synthetic assembled superstructures. Herein, we describe a robust reversible self-assembly system that is composed of a hydrophobic gold nanoparticle (AuNP) core and a shell of pH-responsive dyeincorporated block copolymers. The reversible assembly-disassembly processes were precisely controlled through mediating the molecular interactions between the copolymers and AuNPs. More importantly, the major endogenous biospecies such as proteins will not impair the reversible selfassembly, which was supported by free-energy calculations. The reversible pH-responsive nanostructures were employed as ‘smart’ probes for visualizing the subtle dynamic pH changes among different intracellular compartments, facilitating the study of pH influence on biological processes.

KEYWORDS: surface chemistry; self-assembly; gold nanoparticles; polymers; cellular imaging

Self-assembly chemistry allows the controlled organization of molecular building blocks into a wellconfined nanoarchitecture, which often displays distinct advantages over the molecular counterparts and thus has attracted considerable attention in biomedical applications as diverse as optical sensing and controlled drug delivery.1-6 The molecular self-assembly is driven by various weak and reversible noncovalent forces including electrostatic attraction, π-π stacking, hydrophobic and van der Waals interactions.7 Such weak non-covalent interactions enable the supramolecular systems positively 1

ACS Paragon Plus Environment

ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 30

cooperative in response to subtle perturbation of environmental stimuli.8,9 Thus, stimulus-responsive superstructures can be designed as ‘smart’ probes for sensing dynamic molecular events occurring in complex systems.10-14 pH represents an important physiological parameter that is crucial for maintaining the function of diverse organelles whose pH values are strictly regulated inside cells.15 The abnormal pH variation in organelles could cause the dysfunction of cells, which is linked with many diseases such as cancer and Alzheimer‘s disease.16,17 Therefore, it is highly desired to develop ‘smart’ probes capable of studying the dynamic pH changes in different organelles. Over the past decade, a variety of pH-sensitive small organic dyes and fluorescent proteins have been developed for monitoring intracellular pH and reporting organelle-specific localization,18,19 they however suffer from several hurdles, such as strong background noise, low sensitivity to subtle pH changes, and rapid release from cells. These drawbacks make them difficult to be used for long-term, continuous monitoring of pH-related biological processes occurring inside live cells. Nanotechnology offers great promise in developing pH-responsive nanoparticles.20-22 In particular, self-assembly strategies have been employed to fabricate pH sensors

23-29

capable of

differentiating the acidic compartments such as early endosomes (pH 6.0−6.5) and lysosomes (pH 4.5−5.5). Despite great advances, the polymeric nanoprobes are facing several limitations. First, the reversible assembly-disassembly processes are largely hampered in physiological conditions because the endogeneous biospecies could impair the stability of the polymeric nanoparticles. Second, background fluorescence is still relatively high, due to the limited self-quenching capability of the fluorophores at the off state.24 Third, the polymeric nanoparticles are prepared with a large amount of block copolymers (usually at mg scale), which not only are costly, but also in turn act as potent buffer reagents to impair the endocytic organelles.30 The key of overcoming these issues is to improve the cooperativity of self-assembly superstructures. We herein make use of gold nanoparticles (AuNPs) to prepare a hybrid reversible self-assembly system and modulate the assembly of pH-responsive polymers onto AuNP surfaces through mediating their hydrophobic interactions (Figure 1a). At higher pH values, the neutral polymers prefer to assemble onto the hydrophobic AuNPs; at lower pH values, the pH-responsive tertiary amines (TAs) become protonated and positively charged, thus causing the disassembly of the polymers from the AuNP surfaces. To our surprise, when the acidic solutions were adjusted to be neutral in the presence of proteins, the released polymers prefer to re-assemble onto AuNPs rather than adsorb to the proteins, making it feasible to 2


Page 3 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

modulate the reversible assembly in biological samples. In addition, the AuNP cores serve as highefficiency quenchers (quenching efficiency > 99%) for minimizing background fluorescence,31-35 thus dramatically enhancing the fluorescence activation ratio. Furthermore, only µg-scale block copolymers are required to prepare the colloidal nanoassemblies. Finally, we applied these pH-responsive nanoprobes to report the pH changes of different intracellular compartments, by which the metabolic process of AuNPs can be tracked in live cells.

RESULTS AND DISCUSSION Synthesis and Characterizations of pH-Responsive Self-Assembly Probes. We first synthesized a set of pH-responsive block copolymers by atom-transfer radical polymerization (ATRP) (see details in Supporting Information, Schemes S1-3 and Figures S1-10). These copolymers are comprised of a hydrophilic polyethylene glycol (PEG) chain that functions for maintaining the colloidal dispersity of the nanoassemblies in aqueous solutions36,37 and a hydrophobicity-switchable block bearing TAs with various substituted groups. Three types of TAs including dipentylamine (DPA), diisopropylamine (DiPA), and hexamethyleneimine (HA) were employed to adjust the pKa values (4.0-7.0) of ammonium groups as well as the hydrophobicity of the TA segments. To monitor the reversible pH-responsive property, a pH-insensitive cyanine 5 (Cy5) dye was conjugated to the DiPA-tethered segment as an imaging beacon, producing PEG-b-(PDiPA-r-Cy5). In the neutral Tris-HCl buffer (pH7.4), the DiPA groups were deionized and the segments became hydrophobic. Here, dodecanethiols were added to cover on the AuNPs, making the NP cores hydrophobic, and thus triggering the self-assembly of the copolymers onto the AuNPs. As a result, the typical absorption bands of AuNPs red-shift from 525 to 535 nm in the UV-vis spectra (Supporting Information, Figure S11), along with the appearance of Cy5 band at around 600 nm. The Cy5 dyes were in close proximity to the AuNP surfaces and their fluorescence was effectively quenched (Figure 1b).38 The transmission electron microscopy (TEM) images clearly disclose the core-shell structures of the obtained nanoassemblies with an approximately 3 nm thickness of organic layer surrounding the AuNP cores (Figure 1c, left; Supporting Information, Figure S12b). By measuring the fluorescence intensity of the free polymers before and after assembling on AuNPs, the average amount of copolymers on each 30 nm AuNP was calculated to be 1431 by means of a calibration curve (Supporting Information, Figure S13).

3


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 1. Reversible self-assembly of the pH-responsive block copolymers onto AuNPs. a) Schematic illustration of the preparation of pH-responsive block copolymer-decorated AuNP with the aid of dodecanethiols (purple short lines) and reversible tuning of the self-assembly at different pH values. pHt is referred to as transition pH value. b) Fluorescent spectra of PEG-b-(PDiPA-r-Cy5) (0.05 mg/mL) solutions which were added with AuNPs at various ultimate concentrations: 0, 0.06, 0.12, 0.25, 0.5, and 1.0 nM. c) TEM images of the PEG-b-(PDiPA-r-Cy5)-decorated AuNPs at pH 7.4 and 6.0 solutions, respectively. d) Fluorescent spectra of the PEG-b-(PDiPA-r-Cy5)-decorated AuNPs (0.2 nM) at different pH values: 5.8, 6.2, 6.6, and 7.4. e) Representative fluorescent images of the three PEG-b-(PTA-r-dye)decorated AuNPs at different pH values. f) Plots of normalized fluorescence intensities of the three nanoassemblies versus different pH values. F represents the fluorescence intensity at given pH value, while Fmax and Fmin are the fluorescence intensities in the fully recovered and quenched states, respectively.

4


Page 4 of 30

Page 5 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

We next tested the disassembly behavior of the PEG-b-(PDiPA-r-Cy5)-decorated AuNPs at pH 6.0. Upon adjusting the solution to be acidic, the DiPA groups were positively charged, hence reducing the favorable hydrophobic interactions with the dodecanethiol-coated AuNP cores. Consequently, the copolymers were released from the AuNPs, as detected by a marked increase in fluorescence intensity of Cy5 (Figure 1d). We demonstrated that the fluorescence increase is attributed to the polymers release, rather than the influence of the acidic solution itself because the dyes we used in this study are pH insensitive (Supporting Information, Figure S14). The detachment of the pH-responsive copolymers was also confirmed by the TEM measurement, with clear disappearance of an organic layer on the AuNP cores (Figure 1c, right). It is worth noting that the hydrophobic dodecanethiol-coated AuNPs tend to aggregate in aqueous solutions. However, the TA ionization is a dynamic process, where the unreleased polymers enable the formation of small aggregates (Supporting Information, Figure S15), as verified by a little increase in average hydrodynamic diameters (Supporting Information, Figure S16). Meanwhile, the dodecanethiol-coated AuNPs aggregated slightly on the vial sidewalls, which can be verified by absorbance decrease of the suspending AuNPs in the UV-vis absorption spectra (Supporting Information, Figure S17). Broadening the Self-Assembly Systems with Different pH Transitions. The pH-dependent assembly and disassembly processes were further demonstrated by the DPA- and HA- tethered segments, which were conjugated with tetramethyl rhodamine (TMR) and Cy7.5 dyes, respectively. The physical properties of the three types of copolymers and corresponding nanoassemblies were summarized in Table S1 in Supporting Information. All the three nanoassemblies show sharp fluorescence transitions at different pH values (Figure 1e; Supporting Information, Figure S18). To quantitatively assess the pHresponsive properties, the fluorescence intensities were normalized by the ratio (F-Fmin)/(Fmax-Fmin), where F is the emission fluorescence intensity at desired pH value, while Fmax and Fmin are the emission fluorescence intensities in the fully recovered and quenched states, respectively. The sharpness of the three nanoassemblies was measured by calculating their ΔpH10-90% values, the pH ranges in which the normalized fluorescence intensities vary from 10 to 90%. As indicated in Figure 1f, the ΔpH10-90% values were 0.21, 0.20, and 0.25 pH unit for the DPA-, DiPA-, and HA- tethered nanoassemblies, respectively. Such small values indicate that the nanoassemblies are highly sensitive to slight pH change (about two folds in proton concentration), while that for small organic dyes is nearly 100-fold change since their ΔpH10-90% values are typically 2 pH unit.39,40 In addition, the respective fluorescence transition pH values 5


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(pHt) at which F=(Fmax+Fmin)/2 were determined to be 4.4, 6.2, and 6.8 for the three DPA-, DiPA-, and HA- involved nanoassemblies. Molecular Dynamics Simulations of the Assembly-Disassembly Processes. To investigate the pH-controlled adsorption of the copolymers onto the AuNP surfaces, molecular dynamics (MD) simulations characterizing their spontaneous assembly were performed. To reduce the computational cost, the molecular structure of the copolymers was simplified to be DiPA-conjugated ethylene glycol with three repeating units. As shown in Figure 2a, the copolymers were adsorbed onto AuNP surfaces, with DiPA segments anchoring into the dodecanethiol part at pH > 6.2; while the copolymers were disassembled at pH < 6.2, in line with the experimental observation. To provide a quantitative explanation of the driven force of this pH-controlled adsorption, we made use of free-energy calculation to explore the interactions between a single DiPA segment and an AuNP surface (Figure 2b). As depicted in Figure 2c, the non-protonated DiPA (pH > 6.2) binds much more efficiently to the AuNP surface than the protonated one (pH < 6.2), indicative of a strong attraction between the non-protonated DiPA and the AuNP. Moreover, the free-energy profiles delineating the interactions between two DiPA segments were also calculated. As shown in Figure 2d, a strong repulsive interaction between two protonated DiPA segments is evident. It is apparent that the repulsion between the protonated DiPA segments is much stronger than the attraction between DiPA and AuNP (shown in Figure 2c). The pH-triggered disassembly, therefore, can be ascribed to both the low attraction between the protonated DiPA segments and the hydrophobic AuNP surface, and the high repulsion between the protonated DiPAs in the acidic environment, and vice versa. To delve further into the difference in the interactions between two DiPA segments in the nonprotonated and protonated states, respectively, the electrostatic potential (ESP) of DiPA in different states (Figure 2e) were calculated at the MP2/6-311++G(d, p) level of theory using the Gaussian 09 program.41 DiPA segments become positively charged with the acidification of the solvent, implying a large Columbic repulsive force under such environment, in good agreement with the free-energy calculations.

6


Page 6 of 30

Page 7 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Figure 2. Molecular dynamics simulations for the reversible self-assembly of copolymers onto AuNPs. a) Representative snapshots describing the interactions between the non-protonated and protonated copolymers and the AuNP surfaces. b) Definition of the transition coordinate. , the projection onto the z-axis of the distance between the centers of the mass of DiPA and the AuNP surface extending from 10 to 30 Å, was used to charaterize the the assembling process. d, the distance between the centers of two DiPA segments, was used to characterize their interaction. Free-energy profiles characterize the interactions between c) a DiPA segment and a dodecanethiol-coated AuNP and d) two DiPA segments, respectively. e) Plot of the electrostatic potential for non-protonated (left) and protonated (right) DiPA segments.

Reversibility Evaluation and Free-Energy Calculations. Owing to the reversible ionizable nature of TA groups, the assembly-disassembly processes could be modulated reversibly through switching the pH values of the solutions. To investigate this, the same PEG-b-(PDiPA-r-Cy5)-decorated AuNP conjugates were treated with 0.1 N HCl and NaOH alternatively to enable the ultimate pH values of the aqueous solutions to be 6.0 and 7.4, respectively. The recorded fluorescence spectra indicate two sets of curves corresponding to the recovered and quenched fluorescence emissions at pH 6.0 and 7.4, respectively (Figure 3a). Figure 3b plots the fluorescence intensities of the nanoassemblies in the quenched and recovered states alternatively at least three times. The reversible modulations were supported by the changes of dynamic light scattering (DLS) and UV-vis absorption (Supporting 7


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Information, Figures S19 and S20).

Figure 3. Evaluation of reversibility of the self-assembly system and the possible interference from proteins (BSA). a) Fluorescent spectra of the PEG-b-(PDiPA-r-Cy5)-decorated AuNPs (0.2 nM) during three cycles at pH 7.4 and 6.0 alternatively. b) Fluorescence intensity as a function of repeating pHswitching between pH 7.4 and 6.0. Fluorescence quenching and dequenching in the c) micelle system and d) AuNP-based assembly system at pH 5.0 and pH 6.5 solutions where 10-fold (0.5 mg/mL) BSA are present. e) Schematic illustration for the molecular binding of the copolymer toward AuNP and BSA with different affinity. f) Binding free energies (G) of the copolymer with BSA and AuNP. G=Gsolv+EMM. Gsolv, the solvation free energies, EMM, molecular mechanics energies in the vacuum phase. 8


Page 8 of 30

Page 9 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

We would like to explore if the re-assembly process can be conducted in the presence of proteins that could be a predominant factor impairing the reversibility of the system under physiological conditions. The result for the AuNP-based system was compared with that of the polymeric micelles (see the TEM image of micelles in Supporting Information, Figure S21). Figure 3c indicates that the quenched fluorescence for the micelles prepared with PEG-b-(PDiPA-r-Cy5) (50 µg/mL) can be recovered in pH 5.0 buffer solution with a fluorescence activation ratio of 8.4. When switched the pH value to be 6.5 in the presence of 10-fold bovine serum albumin (BSA, as a model protein, 0.5 mg/mL), > 70% fluorescence remained to be dequenched. The results imply that the presence of proteins can largely affect the re-assembly of copolymers into micellar nanoparticles, which was supported by the DLS measurements (Supporting Information, Figure S22). The hydrodynamic diameter of the assembled micelles at pH 6.5 is around 100 nm, while those decrease dramatically at pH 5.0 due to the disassembly of the micelles. More importantly, when the pH value was swiched back to 6.5, BSA could hamper the assembly of the free copolymers into micelles. In terms of the AuNP-based assembly system that was prepared with the same amount of copolymers as the micelles, 30-fold fluorescence enhancement was achieved upon adjusting the pH value of the colloidal solution from 6.5 to 5.0 (Figure 3d). To our surprise, the addition of 10-fold proteins had negligible influence on the fluorescence quenching when switched the pH value back to 6.5. Note that the presence of 10-fold proteins has no significant interference (P > 0.05) to the fluorescence activation (pH 5.0) in both the micelle and the Au-polymer systems (Supporting Information, Figure S23). We paid attention to the reason why the presence of proteins can scarcely impact the reassembly of the copolymers on AuNPs at pH 6.5. The competitive adsorption of the released copolymers toward proteins and AuNPs was explored using scoring-function-based docking and 600-ns equilibrium simulations.42 As depicted in Figure 3e, both the copolymer-AuNP and copolymer-BSA complexes are thermodynamically stable. To quantitatively compare the binding affinity between the copolymer and AuNP with that between the copolymer and BSA, Molecular Mechanics Poisson-Boltzmann Surface Area (MM-PBSA) approach43 was used to evaluate the binding free energies (G) of the copolymer with AuNP and BSA. As depicted in Figure 3f, the affinity of the copolymer toward AuNP surface is higher than that toward BSA. This difference in binding capability is mainly due to the contribution of the solvation free energy (Gsolv) to the two substrates, where the dodecanethiol-coated AuNPs are more 9


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

hydrophobic than BSA. Therefore, although the interaction between the copolymer and BSA is also thermodynamically favorable, the copolymers prefer to adsorb onto the AuNP surface, in agreement with our experimental results. Analytical Performance Evaluation of the pH-Responsive Probes. To bring these reversible switches into practical applications, we have to evaluate several factors including colloidal stability, responsiveness specificity, and biocompatibility that may impair their analytical performance in real samples. We first explored the colloidal stability of the PEG-b-(PDiPA-r-Cy5)-covered AuNP conjugates in aqueous solutions with various salt concentrations (0.1-1 M). Figure S24 in Supporting Information shows that the fluorescence of the nanoassemblies maintained quenched until the salt concentration reached 1.0 M. Upon mediating the pH value to be 6.0, the fluorescence recovered immediately. The specific responsiveness of these nanoassemblies was investigated in several solutions containing various biospecies including glutathione (GSH), cysteine (Cys), homocysteine (Hcy), human serum albumin (HSA), and hydrogen peroxide (H2O2). Their concentrations were set at the real levels in physiological conditions.44,45 These substances are unable to activate the fluorescence of the nanoassemblies even the solutions were incubated for 24 h (Supporting Information, Figure S25). These results imply that the copolymer-coated AuNP conjugates are highly stable under these extreme conditions. Furthermore, the nanoassemblies show negligible toxicity towards different cell lines even when their concentrations are 2.5 folds higher than that used in the following cell imaging experiments (Supporting Information, Figure S26). Monitoring the Dynamic pH Variation in Different Intracellular Compartments. Encouraged by the above investigations, the pH-responsive nanoassemblies were allowed to report the pH variation in different intracellular compartments. To accelerate the NPs uptake for real-time imaging, the copolymer layers should be tethered with a targeting unit such as Arg-Gly-Asp (RGD) peptide that can bind to αvβ3 integrins expressed on cell surfaces.46 With the aid of RGD, the NPs could be rapidly uptaken via receptor-mediated endocytosis.47 Therefore, we prepared the RGD-terminated copolymers (see Supporting Information, Scheme S4 and Figures S27-30), which were then mixed with the PEG-b(PTA-r-Dye) with a molar ratio of 1:19. The resulting polymer mixtures were grafted onto AuNPs. In this study, we set PEG-b-(PDiPA-r-Cy5)-decorated AuNPs and PEG-b-(PDPA-r-TMR)-decorated AuNPs that were co-assembled with RGD-terminated copolymers as probe (pHt 6.2) and probe (pHt 4.4), respectively. The presence of RGD ligands onto the probes were confirmed by calculating the Au 10


Page 10 of 30

Page 11 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

contents in each HepG2 (human liver carcinoma cell line) cell (Supporting Information, Figure S31). As indicated by the inductively coupled plasma mass spectrometry (ICP-MS) results, 10-fold Au accumulation was observed in the cells treated with RGD-bearing probes (pHt 6.2) over that treated with the same concentration of PEG-b-(PDiPA-r-Cy5)-decorated AuNPs in the absence of RGD. These two probes (0.2 nM for each one) were incubated with HepG2 cells and the imaging results were recorded by confocal microscopy. After entry into cells, the probes were activated, as indicated by the graduate recovery of fluorescence at both Cy5 and TMR channels in 4 h (Figure 4 and Supporting Information, Figure S32 ). With the incubation over time (5-7 h), the fluorescence intensity decreased gradually within the cells. The reversible pH-responsive fluorescence activation/quenching might be due to the reversible assembly of copolymers onto AuNP cores, which were verified by TEM imaging of cell slices (Supporting Information, Figure S33). Before entering into the cells, the NPs are present in close to the cell membrane, and the synthesized polymer shells (2-3 nm thickness) on AuNPs are clear. When the NPs were incubated with the cells for 2-4 h, we can observe that the NPs colocalize with the intracellular compartments (endosomes or lysosomes, which will be validated in the following experiments), where the activatable polymer shells on AuNPs disappear. At 6 h, the AuNPs appear to accumulate in Golgi apparatus with a reassembled 2-3 nm thickness shell around the AuNP cores. Overall, these results indicate that the nanoprobes experienced a typical endocytosis–exocytosis pathway upon interaction with cells;48,49 they first encounter acidic early endosome (pH 6.0–6.5) and then transport to late endosome/lysosome (pH 4.0–5.5), followed by trafficking into the exocytic organelles (pH 7.4) such as Golgi apparatus and multivesicular bodies. To confirm their capability in dynamic imaging, the probes (pHt 6.2) were allowed to monitor the dynamic pH changes mediated by nigericin in live HepG2 cells. Nigericin is a K+/H+ antiporter that is commonly used for modulation of intracellular pH.50 After incubation with probes (pHt 6.2) for 4 h, the adhered cells were washed and further cultured with pH 7.4 buffer containing 10 μM nigericin for different time (0-30 min) (Supporting Information, Figure S34). The results show that the treatment of nigericin leads to efficient fluorescence quenching in 30 min, which was most likely due to the instantaneous acidic-to-neutral pH changes caused by nigericin-mediated high K+/H+ transporting rate. The dynamic self-assembly nanoprobes possess great usefulness for reporting the pH variation in different intracellular compartments.

11


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 4. Confocal microscopy images of a single HepG2 cell that was treated with the mixture of PEGb-(PDiPA-r-Cy5)-decorated AuNPs (refer to as probe (pHt 6.2)) and PEG-b-(PDPA-r-TMR)-decorated AuNPs (refer to as probe (pHt 4.4)) over time (1-7 h). The colloidal concentration of the two probes was identified to be 0.2 nM. The dynamic fluorescence activation of the probe (pHt 6.2) and probe (pHt 4.4) was recorded with Cy5 and TMR channels respectively.

In addition to the single-cell real-time imaging, the time- and pH-dependent activation processes were verified with population cells (10,000 cells) using flow cytometry analysis (Supporting Information, Figure S35). The results revealed that cells treated with the two probes are highly fluorescent at 4 h, with 5-fold intensity higher than that at 2 h. The fluorescence weakened about 2.5 times at 6 h compared to that at 4 h. These flow cytometry results agree well with the confocal imaging data, demonstrating the reversible pH-responsive performance of the probes in cells. Note that each AuNP carries average 1431 copolymers, the concentration of copolymers worked in this study is determined to be 5.6 µg/mL, much lower than that (normally > 50 µg/mL) used in micelle-based nanoprobes.28 Furthermore, when the same amount of free copolymers (e.g., PEG-b-(PDiPA-r-Cy5), 5.6 µg/mL) were incubated with the HepG2 cells, the polymers adsorbed heavily on the cell surfaces or formed aggregates in the medium (Supporting Information, Figure S36), as reflected by the red signals. This is because such low concentration of copolymers can not enable the formation of micelle particles. 12


Page 12 of 30

Page 13 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Finally, we investigated the intracellular transport of AuNPs from early endosomes to late endosome/lysosomes since the two compartments are both acidic while their biological functions are closely associated. We first transfected HepG2 cells with green fluorescent protein (GFP)-fused Rab5a and Lamp1 biomarkers which are able to specifically recognize early endosomes and late endosome/lysosomes, respectively. The GFP-stained cells were incubated with the probes (pHt 6.2) and probes (pHt 4.4) simultaneously, and the colocalization of probes with specific compartments was examined by confocal microscopy imaging at different incubation time. For the probes (pHt 6.2), the Cy5 fluorescence recovered at 2 h, and the signal dots (red) started to overlap with the GFP-labeled early endosomes (green), as demonstrated by the yellow signals in the merge image (Figure 5a). In contrast, no overlapped signals were found with late endosomes/lysosomes at 2 h, indicating that the probes (pHt 6.2) have not reached the late endosomes. Over time, stronger recovered fluorescence of Cy5 was observed, which merged with the GFP signals in both early endosomes and late endosome/lysosomes. The results suggest that some probes (pHt 6.2) still stayed in early endosomes at 4 h, while the others were further activated in late endosome/lysosomes. With further incubation (6 h), the Cy5 fluorescence decreased but no merged signals were observed in the early endosomes, while in the late endosome/lysosomes, some yellow overlapped signals remained to be clearly seen. The results indicate that all the probes (pHt 6.2) have escaped from early endosomes completely in 6 h and some of the probes (pHt 6.2) have entered into the late endosome/lysosomes, while the others have transported into exocytic organelles, as proven by the above TEM experiments. In contrast, the probes (pHt 4.4) remained “silent” in both early endosomes and late endosome/lysosomes at 2 h (Figure 5b). This interesting result might be ascribed to two facts. On one hand, the DPA groups in the probes (pHt 4.4) can not be ionized in early endosome (pH 6.0–6.5), which means that no disassembly occurs; on the other hand, the probes have not reached the late endosomes at 2 h. With 4 h incubation, we can observe the fluorescence recovery from the probes (pHt 4.4), but the signals were only colocalized with late endosomes/lysosomes (pH 4.0–5.5). These results show that the probes (pHt 4.4) were activated exclusively inside the late endosomes/lysosomes owing to their proper pH values. These results imply that, although the two kinds of pH probes can enter the same compartments, their behavior of fluorescence activation is different. This is because the ionization processes of DiPA and DPA in the two probes are strictly governed by the pH values in compartments. As a result, the two probes show different assembly behaviors in these compartments, which could lead 13


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 30

to different metabolism of the nanoparticles. Therefore, we can observe that the activated fluorescence of the two probes is not fully merged throughout the multiple assembly-disassembly processes (Figure 4). In addition, the Pearson's co-localization coefficients (PCC)

51

were allowed to quantify the

colocalization degrees of probe activation signals and endosome/lysosome trackers (Supporting Information, Figure S37). The results further confirmed that the fluorescence recovery of the nanoprobes is strictly dependent on the pH values of the endosomal compartments in cells.

Figure 5. Monitoring the pH-dependent activation of probe (pHt 6.2) and probe (pHt 4.4) in the endosomal compartments of HepG2 cells. The early endosomes and late endosome/lysosomes were labeled with GFP-fused Rab5a and Lamp1 biomarkers, respectively. Representative confocal microscopy images of the GFP-stained HepG2 cells which were separately incubated with a) probe (pHt 6.2) and b) probe (pHt 4.4) for 2, 4, and 6 h. c) The proposed disassembly-reassembly behaviors of probe (pHt 6.2) and probe (pHt 4.4) in different intracellular compartments.

Based on the above investigations, we would like to propose the dynamic disassembly-reassembly processes of the two probes in different intracellular compartments. As described in Figure 5c, after gaining entry into cells, the pH-responsive polymers in probes (pHt 6.2) can be quickly released into the early endosomes, and the disassembly process is saturated as the probes traffic into late 14


Page 15 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

endosome/lysosomes. By contrast, the disassembly of probes (pHt 4.4) takes place in the late endosome/lysosomes exclusively. When the probes are transferred into exocytic organelles such Golgi apparatus, the released polymers are reassembled onto AuNP surfaces, resulting in fluorescence quenching again. This hypothesis was well supported by the single-cell monitoring of fluorescence dynamics, fluorescence colocalization, and TEM analysis.

CONCLUSIONS In summary, we have synthesized a series of pH-responsive copolymers that were reversibly assembled on AuNP cores in aqueous solutions. The assembly and disassembly processes were essentially triggered by tuning the protonation of the TA groups, which allows the reversible manipulation of hydrophobic interactions between the copolymers and the AuNP cores. We have shown here that (i) the reversible assembly-disassembly can be conducted under physiological conditions; (ii) AuNPs act not only as a template for the self-assembly but also as an outstanding quencher for minimizing background fluorescence; (iii) strong fluorescent signals can be activated even though very few copolymers (5.6 µg/mL) anchoring on AuNPs were applied for bioimaging; (iv) the nanoassemblies show extremely sharp pH transitions, indicating high sensitivity to subtle pH changes, and (v) the pHt values of the nanoassemblies can be modulated readily by choosing TAs with various substituted groups. These pH-responsive nanoassemblies were employed as ‘smart’ nanoprobes to report the dynamic pH changes between different intracellular compartments, which facilitates tracking the transport processes of NPs in living cells. Undoubtedly, the strategy to reversibly control surface chemistry can be applied to manipulate the properties of other types of nanoparticles, greatly extending the use of the “smart” responsive platforms in a variety of biological and biomedical fields.

MATERIALS AND METHODS General Materials 6-Carboxytetramethylrhodamine N-succinimidyl ester (6-TAMRA-SE) was purchased from the Lumiprobe Corporation. MeO-PEG114-OH, (2-hexamethylene imino) ethanol (HA), and chloroauric acid were purchased from Sigma-Aldrich. 2-(diisopropyl amino)ethyl methacrylate (DiPA-MA), bromopentane, ethanolamine, methacryloyl chloride, sodium salts and 1-dodecanethiol were purchased from Alfa Aesar Company. 2-aminoethyl methacrylate (AMA) was purchased from J&K Chemicals and 15


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

it was recrystallized twice with isopropanol and ethyl acetate (3:7) before use. Other solvents and reagents were used as received from Sigma-Aldrich or Alfa Aesar Company.

Synthesis of PEG Macroinitiator. PEG114-OH (5 g, 0.001 mol) was dissolved in 20 mL of CH2Cl2 in a 100 mL three-necked, round-bottomed flask. Triethylamine (0.29 mL, 0.002 mol) was added and the solution was cooled to 0 oC. Then, 2-bromoisobutyryl bromide (0.25 mL, 0.002 mol) was distributed in 5 mL CH2Cl2 and the mixture was added dropwise into the flask by constant pressure funnel over 1 h. The reaction was stirred at room temperature for another 6 h. The resulting mixture was filtration, washed with saturated brine and NaHCO3 respectively, dried over MgSO4. The solid residue was removed by filtration, then the filtrate was concentrated and precipitation into cold ether and dried in vacuum to obtain a white solid product. Yield: 80%. The 1H NMR spectrum for the product is shown in Figure S1.

Synthesis of 2-(Dipentylamino) Ethanol. First, we added ethanolamine (3.1 g, 0.05 mol) in 100 mL acetonitrile under mild stirring. After ethanolamine was dissolved, bromopentane (15.1 g, 0.1 mol) and Na2CO3 (13.25 g, 0.125 mol) were added to the solution. After stirring at room temperature for 6 h, the solution was filtered to remove the insoluble substance and the solvent was evaporated. The crude product was purified by column chromatography (CH2Cl2:CH3OH=10:1) with trace ammonia to obtain 2-(dipentylamino) ethanol as a light yellow liquid. Yield: 75%. The 1H NMR spectrum for the product is shown in Figure S2.

Synthesis of Methacrylate Monomers. We synthesized 2-(dipentylamino) ethyl methacrylate (DPAMA) and 2-(hexamethylene imino) ethyl methacrylate (HA-MA) using a similar procedure. Here we described the synthetic method for DPA-MA as an example. Hydroquinone (0.22 g, 0.002 mol), 2(dipentylamino) ethanol (48.6 g, 0.2 mol), and triethylamine (20.2 g, 0.2 mol) were dissolved in 200 mL THF. The mixture was incubated in ice-salt bath for 15 min, then methacryloyl chloride (20.8 g, 0.2 mol) was added dropwise into the flask slowly. The solution was refluxed for 4 h. Afterwards the mixture was filtered and concentrated to dryness. The residual product was purified by reduced pressure distillation and afforded a light yellow liquid. The yields for DPA-MA and HA-MA are calculated to be 80% and 60%, respectively. The 1H NMR spectra of the obtained DPA-MA and HA-MA are shown in Figures S3 and S6, respectively. 16


Page 16 of 30

Page 17 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Synthesis of Dye-Conjugated Block Copolymers. We synthesized PEG-b-tertiary amine (TA) copolymers (refer to as PEG-b-(PTA-r-AMA)) using a living radical polymerization method named atom transfer radical polymerization (ATRP). The synthetic route for PEG-b-(PDPA-r-AMA) is used as a general sample. All the monomers, catalyst and initiator including 2-aminoethyl methacrylate (AMA, 25 mg, 0.15 mmol) 2- (dipentylamino) ethyl methacrylate (DPA-MA, 336 mg, 1.25 mmol), PMDETA (5.5 ml, 0.025 mmol), and PEG114-Br (125mg, 0.025 mmol) were added into a mixed solvent of 2-propanol (1 mL) and DMF (1 mL). The reaction tube was degassed by three vacuum/argon cycles. Then the purified white powder of CuBr (4 mg, 0.025 mmol) was quickly added into the tube, after that another three vacuum/argon cycles were carried out. The mixture was maintained at 60 oC for 12 h. The crude product was then diluted with 20 mL THF. Copper catalyst was removed by passing a solution of copolymer in THF through a column of basic alumina. The purified copolymer was recovered by dialyzed in distilled water and lyophilized to obtain a white powder. The molecular weights (Mn and Mw) and polydispersity index (PDI) of each copolymer were listed in Table S1. The gel permeation chromatography system (GPC, Waters, USA) was used to measure the molecular weights and their corresponding distribution of polymers. THF was used as the eluent at a low flow rate of 1.0 mL/min. Monodispersed polystyrene standards were used to create the calibration curves. The yields for PEG-b(PDPA-r-AMA), PEG-b-(PDiPA-r-AMA), and PEG-b-(PHA-r-AMA) are calculated to be 45%, 64%, and 55%, and their corresponding 1H NMR spectra are shown in Figures S4, S5, and S7, respectively. We conjugated dyes with the copolymers in the same step. In a representative procedure, PEG-b(PDPA-r-AMA) (20 mg) was dissolved in 2 mL DMF, then tetramethyl rhodamine (TMR) with NHS ester was added (molar ratio of dye to tertiary amine 1.5:1). The reaction was carried out in the dark at room temperature for two days. After that the mixture was purified by Sephadex LH-20 (CH3OH:CHCl3=1:1) to remove the unreacted fluorescent dyes. The produced copolymers were lyophilized and dissolved in DMF as a stock solution at 4 oC for storage.

Synthesis of RGD-Terminated Copolymers. Arg-Gly-Asp (RGD) peptides (cyclo (Arg-Gly-Asp-DTyr-Lys (CO-CH2SH)), cRGD-SH) were purchased from GL Biochem (Shanghai) Ltd. The peptides were

tethered

on

the

nanoprobes

by

introducing

maleimide-terminated

PEG-b-poly

(2-

(Pentamethyleneimino) ethyl methacrylate) (MAL-PEG-b-PPEA) (see Supporting Information, Scheme 17


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

S4). First, we added the mixture of furan (4.08g, 60 mmol) and N-Maleoyl-β-alanine 1 (1.01 g, 6 mmol) into 20 mL benzene at room temperature. The mixture was heated to 80 oC and refluxed for 8 h. After that, the solution was cooled to room temperature to produce a white precipitate. The solvent was removed by concentrated in vacuo. The obtained residue was then purified by column chromatography (CH2Cl2:CH3OH=3:1) to yield product 2 as a white powder. Yield: 80 %. N,N′-dicyclohexylcarbodiimide (200 mg, 0.96 mmol) and trace amount of dimethylaminopyridine was added to a mixture of N-hydroxymaleimide (100 mg, 0.88 mmol) and product 2 (192 mg, 0.8 mmol) in 100 mL dichloromethane. The solution was stirred at 0 oC for 20 min and transferred to room temperature for another 2 h. HO-PEG5000-NH2 (2 g, 0.4 mmol) was added to the mixture drop by drop, then the reaction system was stirred overnight. The reaction solution was filtered and concentrated to 10 mL byrotary evaporation. The concentrated liquor was poured into 150 mL cold diethyl ether and the precipitate was collected and dried to yield product proMAL-PEG5000. Yield: 59%. Triethylamine (280 µL, 2 mmol) was added in 20 mL dichloromethane and mixed with proMALPEG5000. Then, 2-Bromo-2-methylpropionyl bromide (260 µL, 2 mmol) was added slowly and the solution was stirred overnight under argon protection. The resulted solution was poured into 200 mL cold diethyl ether and the residual was collected. Two more dissolve-and-precipitation procedure of the crude product of PEG macroinitiator was repeated and then the product was dried. All the monomer, catalyst and initiator including PEA (276 mg, 1.4 mmol), PMDETA (7.0 µL, 0.034 mmol), and proMALPEG5000-Br (140 mg, 0.028mmol) were added in a mixed solvent of 2-propanol (1 mL) and DMF (1 mL). The reaction tube was degassed by three vacuum/argon cycles. Then, purified white powder of CuBr (4.5mg, 0.031mmol) was quickly added into the tube, after that another three vacuum/argon cycles were carried out. Polymerization was reacted at 60 oC for 12 h. The crude product was then diluted with 20 mL THF. Copper catalyst was removed by passing a solution of copolymer in THF through a column of basic alumina. The purified copolymer was recovered by dialyzed in distilled water and lyophilized to obtain proMAL-PEG-b-PPEA as a white powder. Yield: 79%. To deprotect the copolymer, 100 mg proMAL-PEG-b-PPEA copolymer in 10 mL toluene was refluxed for 8 h. Then the solvent was removed by rotary evaporation. The purified copolymer was dialyzed in distilled water and lyophilized to obtain MAL-PEG-b-PPEA as a white powder. Yield: 89%. All the products were characterized by 1H NMR, as shown in Figures S23-26. For RGD conjugation, MAL-PEG-PPEA was dissolved in THF and the solution was added into 18


Page 18 of 30

Page 19 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

distilled water under sonication. Then the hybrid was ultra-filtered at 10000 rpm for 10 minutes to remove THF. Then three times excess cRGD peptides were added, and the mixture was shaken at room temperature for 4 h. Finally, the mixture was ultra-filtered four times to remove free cRGD residues. The crude products were freeze-dried to yield a white powder.

Synthesis of 30 nm AuNPs. All glass containers and the magnetic stir-bars were immersed in aqua regia for 30 min and washed thoroughly with deionized water. 13 nm gold nanoparticles (AuNPs) were synthesized by sodium citrate reduction of HAuCl4. Briefly, to a 200 mL boiling aqueous solution of HAuCl4 (82 mg, 2 mmol) was added trisodium citrate (236 mg, 80 mmol) rapidly, then the solution was heated with vigorous stirring for additional 20 min. Subsequently, the resulting solution was cooled to room temperature. 30 nm AuNPs were prepared according to the seed growth method via reduction of HAuCl4 with NH2OH.52 To a 250 mL aqueous solution of 13 nm AuNPs (0.1 nmol) was added NH2OH (0.2 mol, 2 mL) rapidly. After 5 min stirring at room temperature, 208 μL of HAuCl4 (100 mM) was added dropwise for 10 min, the mixture turned red gradually and stirred for another 30 min, 5% trisodium citrate (80 mg, 2 mL) was added to stabilize the 30 nm AuNPs.

Coating AuNPs with Copolymers. Since the excess citrate ions in solution could influence the particle encapsulation, the as-prepared citrate-stabilized AuNPs were centrifuged and redispersed in water prior to dispersion in DMF. First, 1 mL aqueous solution of AuNPs solution (30 nm, 0.4 nM) was centrifuged at 4,000 × g for 10 min. After removing the supernatants, the concentrated AuNPs were redispersed and then slowly dripped with 1 mL of DMF. The DMF suspensions were prepared freshly before use. Next, 100 µL PEG-b-(PTA-r-Dye) copolymers (1 mg/mL) in DMF was added to 900 µL AuNPs solution (30 nm, 0.4 nM in DMF) with vigorous stirring. Then, 0.25 mL of Tris-HCl (pH 7.4) was added dropwise (0.5 mL/ h) via micro-injection pump at room temperature. After that 20 µL dodecanethiol solution (10 L in 1 mL DMF) was added to the solution and stirred for 1 h. Another 3 mL of Tris-HCl (pH 7.4) was added dropwise (2 mL/ h) to the mixture by micro-injection pump. Organic solvent was removed by dialysis in PBS (pH 7.4, 10 mM) for 12 h. Finally, the solution in the dialysis bag was transferred to centrifuge tube and was then centrifuged at 4,000 × g for 30 min. This procedure was repeated four times to remove the free polymers. The purified copolymer-decorated AuNPs were redispersed in 1 mL PBS (pH 7.4). These procedures are amenable to the RGD-conjugated nanoprobes. 19


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

The polymers used for encapsulating AuNPs are a mixture of RGD-PEG-PPEA and PEG-b-(PTA-r-Dye) with a molar ratio of 1:19.

Recording Fluorescence Spectra. The fluorescence spectra of PEG-b-(PTA-r-Dye)-decorated AuNPs were obtained on a Hitachi fluorometer (F-7500 model). The nanoconjugates in high concentration were diluted to desired concentrations by 0.2 M citric-phosphate buffers with different pH values. The colloidal concentration for the fluorescence measurement in solutions was fixed to be 0.2 nM. The PEGb-(PDiPA-r-Cy5)-decorated AuNPs were excited at 633 nm, and the emission spectra was collected from 650 to 700 nm. The PEG-b-(PDPA-r-TMR)-decorated AuNPs were excited at 530 nm, and the emission spectra was collected from 550 to 650 nm. The PEG-b-(PHA-r-Cy7.5)-decorated AuNPs were excited at 780 nm, and the emission spectra was collected from 800 to 900 nm. Both the emission and excitation slits were 5 nm. The emission intensity at peak value was used to quantify the pH responsiveness for the nanoconjugates.

pH Titration and pKa Measurement. The pH titration of the three types of PEG-b-(PTA-r-Dye) copolymers was conducted following a similar method. For example, PEG-b-(PDA-r-TMR) (27.5 mg) was added with HCl (0.1 mol, 5 mL) in a small beaker. After the copolymers were totally dissolved, pH titration was carried out by adding 100 L aliquots of 0.1 mol NaOH solution under constant stirring. Then, the pH values were measured using a Thermal pH meter. The pKa values were calculated as the pH in the middle of the two equivalence points in the titration curve. The pKa values for all the three types of PEG-b-(PTA-r-Dye) copolymers were listed in Table S1.

Cell Culture. HepG2 (hepatocellular carcinoma cell lines) and 3T3 (fibroblast cell lines) cells were cultured in a DMEM medium (Roswell Park Memorial Institute medium) supplemented with 10 % fetal bovine serum (FBS) and 1 % penicillin-streptomycin, respectively. All the cells were placed in a humidified incubator at 37 oC with 5 % CO2 until they were confluent. Before use, cells were grown to confluence, washed with PBS, desorbed from culture plates with trypsin (GIBCO) and resuspended in corresponding fresh culture medium.

Cell Transfection. For colocalization experiment, HepG2 cells were transfected with GFP-Rab5a and 20


Page 20 of 30

Page 21 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

GFP-Lamp1 plasmids that were both encapsulated by lipo2000. Rab5a and Lamp1 plasmids are endosome marker and lysosome marker, respectively. One days before transfection, HepG2 cells were inoculated in confocal dish to ensure that the cell confluence reached 70% when transfected. The plasmids were diluted with 25 μL Opti-MEM. Then the lipo2000 was mixed with the Opti-MEM solution, gently homogenized, and incubated at room temperature for 20 min. The composite was laid to the confocal dish, and the plates were swayed back and forth to make the compounds distribute evenly.

Cytotoxicity Evaluation. The cytotoxicity of PEG-b-(PTA-r-Dye)-decorated AuNPs was evaluated using MTT assays in a 96-well plate (a density of 5,000 cells per well). Both the HepG2 and 3T3 fibroblast cell lines were independently incubated with increasing concentrations (from 0.1 to 0.5 nM) of nanoprobes which dispersed in culture medium with 10% FBS for 24 h in an incubator (37 C, 5% CO2). After that, a 10 µL MTT (5 mg/mL) was added to each well and incubated with all the cells for additional 3 h in the incubator. Ultimately, the formazan products were measured by using microplate reader at the absorbance of 490 nm.

Real-Time Confocal Imaging and Colocalization. We seeded HepG2 cells in confocal dishes with 2 mL complete DMEM medium and cultured for 24 h. PEG-b-(PDiPA-r-Cy5)-decorated AuNPs (refer to as probe pHt 6.2) and PEG-b-(PHA-r-Cy7.5)-decorated AuNPs (refer to as probe pHt 4.4) that were coassembled with RGD peptides were dissolved in culture medium with a final concentration of 0.2 nM, respectively. The probes were incubated with the cells for 1 h, and the free probes in the medium were removed. The confocal images of individual cell were captured at 1, 2, 3, 4, 5, 6, and 7 h. For cellular colocalization experiments, after 24 h of cell transfection, the probes were incubated with the cells for 2, 4, and 6 h. The stained cells were washed with PBS for 5 times, fixed with paraformaldehyde for 20 min, and washed with PBS for 3 times once again. The confocal images were captured by a Leica TCS-SP8 confocal microscope with a 40 × objective oil lens. GFP, TMR and Cy5 were excited at 488, 530 and 633 nm, respectively.

Flow Cytometry. HepG2 Cells were inoculated on 6 well plates and cultured overnight to grown confluence. The PEG-b-(PDiPA-r-Cy5)-decorated AuNPs and PEG-b-(PHA-r-Cy5)-decorated AuNPs were cultured with the cells for 2, 4, and 6 h, respectively. Then the cells were washed with PBS and 21


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

treated with trypsin to detach from the well plates. The cell solution was centrifuged (1,000 g, 3 min), then resuspended and washed three times with PBS. After fixation by paraformaldehyde (4%), the cells were washed with PBS for three times. BD LSR Fortessa was used in the flow cytometry experiment with excitation at 640 nm. 10,000 cells were used in the measurement of each sample.

Molecular Dynamics Simulations. All the MD simulations described herein were conducted using the program NAMD 2.1253 with the CHARMM General Force Field 54 and the TIP3P water model.55 The temperature and the pressure were maintained at 300 K and 1 atm, respectively, employing Langevin dynamics56 and Langevin piston method.57 The length of the covalent bonds involving a hydrogen atom was frozen to its equilibrium value by means of the SHAKE/RATTLE58,59 and SETTLE algorithms.60 The r-RESPA multiple time-step algorithm61 was utilized to integrate the equations of motion with a time step of 2 and 4 fs for short- and long-range interactions, respectively. Long-range electrostatic forces were evaluated using the particle mesh Ewald approach.62 A smoothed 12 Å spherical cutoff was used to truncate van der Waals and short-range Columbic interactions. Visualization and analysis of MD trajectories were carried out with the VMD 1.9.2 package.63 Molecular docking program AutoDock was employed for a systematic search of the possible binding sites of polymer with respect to the protein BSA.

Free-Energy Calculations. All the PMF calculations were carried out using the meta-eABF method.64 The transition coordinates of the two free-energy calculations were chosen as , the distance between a TA segment and a dodecanethiol-coated AuNP and d, the distance between two TA segments, respectively. The time for the simulation of each molecular assembly was set to be 500 ns.

PCC calculation. PCC is one of the standard statistical techniques in identifying the matching degree between two signals. To calculate the PCC of a pair of fluorescent images, all pixels having the same image coordinates are paired, such as green pixels and red pixels. PCC generates a range of values from 1, a perfect positive correlation, to -1, a perfect but inverse correlation, with 0 representing a random distribution. PCC is calculated according to the following equation,51 𝑃𝐶𝐶 =

∑(𝑅𝑖 ― 𝑅𝑎𝑣𝑒𝑟) ∙ (𝐺𝑖 ― 𝐺𝑎𝑣𝑒𝑟) ∑(𝑅𝑖 ― 𝑅𝑎𝑣𝑒𝑟)2 ∙ ∑(𝐺𝑖 ― 𝐺𝑎𝑣𝑒𝑟)2 22


Page 22 of 30

Page 23 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Ri: intensity in red channel; Raver: average intensity in red channel; Gi: intensity in green channel; Gaver: average intensity in green channel.

ASSOCIATED CONTENT Supporting Information Supplementary Schemes S1-4, Table S1, and Figures S1-37 are included in the Supporting Information. This material is available free of charge via the internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author *[email protected].

ACKNOWLEDGMENTS We acknowledge the support from the National Natural Science Foundation of China (21775075), the Fundamental Research Funds for Central Universities (China), and the Thousand Youth Talents Plan of China.

REFERENCES AND NOTES 1.

Datta, S.; Saha, M. L.; Stang, P. J. Hierarchical Assemblies of Supramolecular Coordination Complexes. Acc. Chem. Res. 2018, 51, 2047−2063.

2.

Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. A DNA-Based Method for Rationally Assembling Nanoparticles into Macroscopic Materials. Nature 1996, 382, 607−609.

3.

Whitesides, G. M.; Grzybowski, B. Self-Assembly at All Scales. Science 2002, 295, 2481−2421.

4.

Douglas, S. M.; Dietz, H.; Liedl, T.; Hogberg, B.; Graf, F.; Shih, W. M. Self-Assembly of DNA into Nanoscale Three-Dimensional Shapes. Nature 2009, 459, 414−418.

5.

Li, Y.; Wang, Y.; Huang, G.; Gao, J. Cooperativity Principles in Self-Assembled Nanomedicine. Chem. Rev. 2018, 118, 5359−5391.

6.

Song, J.; Zhou, J.; Duan, H. Self-Assembled Plasmonic Vesicles of SERS-Encoded Amphiphilic Gold Nanoparticles for Cancer Cell Targeting and Traceable Intracellular Drug Delivery. J. Am. Chem. Soc. 2012, 134, 13458−13469. 23


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

7.

Xing, P.; Zhao, Y. Multifunctional Nanoparticles Self-Assembled from Small Organic Building Blocks for Biomedicine. Adv. Mater. 2016, 28, 7304–7339.

8.

Whitesides, G.; Mathias, J.; Seto, C. Molecular Self-Assembly and Nanochemistry: a Chemical Strategy for the Synthesis of Nanostructures. Science 1991, 254, 1312−1319.

9.

Rosi, N. L.; Mirkin, C. A. Nanostructures in Biodiagnostics. Chem. Rev. 2005, 105, 1547−1562.

10. Seferos, D. S.; Giljohann, D. A.; Hill, H. D.; Prigodich, A. E.; Mirkin, C. A. Nano-Flares: Probes for Transfection and mRNA Detection in Living Cells. J. Am. Chem. Soc. 2007, 129, 15477−15479. 11. Prigodich, A. E.; Randeria, P. S.; Briley, W. E.; Kim, N. J.; Daniel, W. L.; Giljohann, D. A.; Mirkin, C. A. Multiplexed Nanoflares: mRNA Detection in Live Cells. Anal. Chem. 2012, 84, 2062−2066. 12. Ye, D.; Shuhendler, A. J.; Cui, L.; Tong, L.; Tee, S. S.; Tikhomirov, G. A.; Felsher, D. W.; Rao, J. Bioorthogonal Cyclization-Mediated in situ Self-Assembly of Small-Molecule Probes for Imaging Caspase Activity in vivo. Nat. Chem. 2014, 6, 519−526. 13. Xu, L. G.; Gao, Y. F.; Kuang, H.; Liz-Marzan, L. M.; Xu, C. L. MicroRNA-Directed Intracellular Self-Assembly of Chiral Nanorod Dimers. Angew. Chem., Int. Ed. 2018, 57, 10544−10548. 14. Wu, Z.; Liu, G. Q.; Yang, X. L.; Jiang, J. H. Electrostatic Nucleic Acid Nanoassembly Enables Hybridization Chain Reaction in Living Cells for Ultrasensitive mRNA Imaging. J. Am. Chem. Soc. 2015, 137, 6829−6836. 15. Llopis, J.; McCaffery, J. M.; Miyawaki, A.; Farquhar, M. G.; Tsien, R. Y. Measurement of Cytosolic, Mitochondrial, and Golgi pH in Single Living Cells with Green Fluorescent Proteins. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 6803−6808. 16. Webb, B. A.; Chimenti, M.; Jacobson, M. P.; Barber, D. L. Dysregulated pH: a Perfect Storm for Cancer Progression. Nat. Rev. Cancer 2011, 11, 671−677. 17. Fang, B.; Wang, D.; Huang, M.; Yu, G.; Li, H. Hypothesis on the Relationship between the Change in Intracellular pH and Incidence of Sporadic Alzheimer's Disease or Vascular Dementia. Int. J. Neurosci. 2010, 120, 591–595. 18. Lee, M. H.; Han, J. H.; Lee, J. H.; Park, N.; Kumar, R.; Kang, C.; Kim, J. S. Two-Color Probe to Monitor a Wide Range of pH Values in Cells. Angew. Chem., Int. Ed. 2013, 52, 6206–6209. 19. Wu, L.; Wang, Y.; James, T. D.; Jia, N.; Huang, C. A Hemicyanine Based Ratiometric Fluorescence Probe for Mapping Lysosomal pH during Heat Stroke in Living Cells. Chem. Commun. 2018, 54, 5518–5521. 24


Page 24 of 30

Page 25 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

20. Marín, M. J.; Galindo, F.; Thomas, P.; Russell, D. A. Localized Intracellular pH Measurement Using a Ratiometric Photoinduced Electron-Transfer-Based Nanosensor. Angew. Chem., Int. Ed. 2012, 51, 9657–9661. 21. Dennis, A. M.; Rhee, W. J.; Sotto, D.; Dublin, S. N.; Bao, G. Quantum Dot-Fluorescent Protein FRET Probes for Sensing Intracellular pH. ACS Nano 2012, 6, 2917–2924. 22. Shi, W.; Li, X.; Ma, H. A Tunable Ratiometric pH Sensor Based on Carbon Nanodots for the Quantitative Measurement of the Intracellular pH of Whole Cells. Angew. Chem., Int. Ed. 2012, 51, 6432–6435. 23. Zhou, K.; Wang, Y.; Huang, X.; Luby-Phelps, K.; Sumer, B. D.; Gao, J. Tunable, Ultrasensitive pHResponsive Nanoparticles Targeting Specific Endocytic Organelles in Living Cells. Angew. Chem., Int. Ed. 2011, 50, 6109−6114. 24. Ma, X.; Wang, Y.; Zhao, T.; Li, Y.; Su, L.-C.; Wang, Z.; Huang, G.; Sumer, B. D.; Gao, J. UltrapH-Sensitive Nanoprobe Library with Broad pH Tunability and Fluorescence Emissions. J. Am. Chem. Soc. 2014, 136, 11085−11092. 25. Wang, Y.; Zhou, K.; Huang, G.; Hensley, C.; Huang, X.; Ma, X.; Zhao, T.; Sumer, B. D.; DeBerardinis, R. J.; Gao, J. A Nanoparticle-Based Strategy for the Imaging of a Broad Range of Tumours by Nonlinear Amplification of Microenvironment Signals. Nat. Mater. 2013, 13, 204−212. 26. Wang, C.; Zhao, T.; Li, Y.; Huang, G.; White, M. A.; Gao, J. Investigation of Endosome and Lysosome Biology by Ultra pH-Sensitive Nanoprobes. Adv. Drug Delivery Rev. 2017, 113, 87−96. 27. Li, Y.; Zhao, T.; Wang, C.; Lin, Z.; Huang, G.; Sumer, B. D.; Gao, J. Molecular Basis of Cooperativity in pH-Triggered Supramolecular Self-Assembly. Nat. Commun. 2016, 7, 13214. 28. Wang, Y.; Wang, C.; Li, Y.; Huang, G.; Zhao, T.; Ma, X.; Wang, Z.; Sumer, B. D.; White, M. A.; Gao, J. Digitization of Endocytic pH by Hybrid Ultra-pH-Sensitive Nanoprobes at Single-Organelle Resolution. Adv. Mater. 2017, 29, 1603794. 29. Zhao, T.; Huang, G.; Li, Y.; Yang, S.; Ramezani, S.; Lin, Z.; Wang, Y.; Ma, X.; Zeng, Z.; Luo, M.; de Boer, E.; Xie, X.-J.; Thibodeaux, J.; Brekken, R. A.; Sun, X.; Sumer, B. D.; Gao, J. A TransistorLike pH Nanoprobe for Tumour Detection and Image-Guided Surgery. Nat. Biomed. Eng. 2017, 1, 0006. 30. Wang, C.; Wang, Y.; Li, Y.; Bodemann, B.; Zhao, T.; Ma, X.; Huang, G.; Hu, Z.; DeBerardinis, R. J.; White, M. A.; Gao, J. A Nanobuffer Reporter Library for Fine-Scale Imaging and Perturbation 25


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

of Endocytic Organelles. Nat. Commun. 2015, 6, 8524. 31. Dulkeith, E.; Morteani, A. C.; Niedereichholz, T.; Klar, T. A.; Feldmann, J.; Levi, S. A.; van Veggel, F. C. J. M.; Reinhoudt, D. N.; Moller, M.; Gittins, D. I. Fluorescence Quenching of Dye Molecules Near Gold Nanoparticles: Radiative and Nonradiative Effects. Phys. Rev. Lett. 2002, 89, 203002. 32. You, C.-C.; Miranda, O. R.; Gider, B.; Ghosh, P. S.; Kim, I.-B.; Erdogan, B.; Krovi, S. A.; Bunz, U. H. F.; Rotello, V. M. Detection and Identification of Proteins Using Nanoparticle-Fluorescent Polymer 'Chemical Nose' Sensors. Nat. Nanotechnol. 2007, 2, 318−323. 33. Bajaj, A.; Miranda, O. R.; Kim, I. B.; Phillips, R. L.; Jerry, D. J.; Bunz, U. H. F.; Rotello, V. M. Detection and Differentiation of Normal, Cancerous, and Metastatic Cells Using NanoparticlePolymer Sensor Arrays. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 10912−10916. 34. De, M.; Rana, S.; Akpinar, H.; Miranda, O. R.; Arvizo, R. R.; Bunz, U. H. F.; Rotello, V. M. Sensing of Proteins in Human Serum Using Conjugates of Nanoparticles and Green Fluorescent Protein. Nat. Chem. 2009, 1, 461−465. 35. Rana, S.; Le, N. D. B.; Mout, R.; Saha, K.; Tonga, G. Y.; Bain, R. E. S.; Miranda, O. R.; Rotello, C. M.; Rotello, V. M. A Multichannel Nanosensor for Instantaneous Readout of Cancer Drug Mechanisms. Nat. Nanotechnol. 2015, 10, 65−69. 36. Walkey, C. D.; Olsen, J. B.; Guo, H.; Emili, A.; Chan, W. C. W. Nanoparticle Size and Surface Chemistry Determine Serum Protein Adsorption and Macrophage Uptake. J. Am. Chem. Soc. 2012, 134, 2139−2147. 37. Dai, Q.; Walkey, C.; Chan, W. C. W. Polyethylene Glycol Backfilling Mitigates the Negative Impact of the Protein Corona on Nanoparticle Cell Targeting. Angew. Chem., Int. Ed. 2014, 53, 5093−5096. 38. Li, N.; Chang, C.; Pan, W.; Tang, B. A Multicolor Nanoprobe for Detection and Imaging of TumorRelated mRNAs in Living Cells. Angew. Chem., Int. Ed. 2012, 51, 7426−7430. 39. Dong, Z.; Han, Q.; Mou, Z.; Li, G.; Liu, W. A Reversible Frequency Upconversion Probe for RealTime Intracellular Lysosome-pH Detection and Subcellular Imaging. J. Mater. Chem. B 2018, 6, 1322−1327. 40. Yin, J.; Hu, Y.; Yoon, J. Fluorescent Probes and Bioimaging: Alkali Metals, Alkaline Earth Metals and pH. Chem. Soc. Rev. 2015, 44, 4619−4644. 41. Frisch, M. J.; Trucks, G. W.; Schlegel, H. B.; Scuseria, G. E.; Robb, M. A.; Cheeseman, J. R.; Scalmani, G.; Barone, V.; Mennucci, B.; Petersson, G. A.; Nakatsuji, H.; Caricato, M.; Li, X.; 26


Page 26 of 30

Page 27 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Hratchian, H. P.; Izmaylov, A. F.; Bloino, J.; Zheng, G.; Sonnenberg, J. L.; Hada, M.; Ehara, M.; et al. Gaussian 09, Revision D1; Gaussian, Inc.: Wallingford, CT. 2009. 42. Meng, E. C.; Shoichet, B. K.; Kuntz, I. D. Automated Docking with Grid-Based Energy Evaluation. J. Comput. Chem. 1992, 13, 505−524. 43. Kollman, P. A.; Massova, I.; Reyes, C.; Kuhn, B.; Huo, S.; Chong, L.; Lee, M.; Lee, T.; Duan, Y.; Wang, W.; Donini, O.; Cieplak, P.; Srini-vasan, J.; Case, D. A.; Cheatham, T. E. Calculating Structures and Free Energies of Complex Molecules: Combining Molecular Mechanics and Continuum Models. Acc. Chem. Res. 2000, 33, 889−897. 44. Turell, L.; Radi, R.; Alvarez, B. The Thiol Pool in Human Plasma: the Central Contribution of Albumin to Redox Processes. Free Radical Biol. Med. 2013, 65, 244−253. 45. Miller, E. W.; Albers, A. E.; Pralle, A.; Isacoff, E. Y.; Chang, C. J. Boronate-Based Fluorescent Probes for Imaging Cellular Hydrogen Peroxide. J. Am. Chem. Soc. 2005, 127, 16652−16659. 46. Majzoub, R. N.; Chan, C.-L.; Ewert, K. K.; Silva, B. F. B.; Liang, K. S.; Jacovetty, E. L.; Carragher, B.; Potter, C. S.; Safinya, C. R. Uptake and Transfection Efficiency of PEGylated Cationic Liposome-DNA Complexes with and without RGD-Tagging. Biomaterials 2014, 35, 4996−5005. 47. Almutairi, A.; Rossin, R.; Shokeen, M.; Hagooly, A.; Ananth, A.; Capoccia, B.; Guillaudeu, S.; Abendschein, D.; Anderson, C. J.; Welch, M. J.; Fréchet, J. M. J. Biodegradable Dendritic PositronEmitting Nanoprobes for the Noninvasive Imaging of Angiogenesis. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 685–690. 48. Xu, R.; Huang, L.; Wei, W.; Chen, X.; Zhang, X.; Zhang, X. Real-Time Imaging and Tracking of Ultrastable Organic Dye Nanoparticles in Living Cells. Biomaterials 2016, 93, 38–47. 49. Jiang, X.; Röcker, C.; Hafner, M.; Brandholt, S.; Dörlich, R. M.; Nienhaus, G. U. Endo- and Exocytosis of Zwitterionic Quantum Dot Nanoparticles by Live HeLa Cells. ACS Nano 2010, 4, 6787–6797. 50. Lee, C.; Kang, H. J.; von Ballmoos, C.; Newstead, S.; Uzdavinys, P.; Dotson, D. L.; Iwata, S.; Beckstein, O.; Cameron, A. D.; Drew, D. A Two-Domain Elevator Mechanism for Sodium/Proton Antiport. Nature 2013, 501, 573−577. 51. Barlow, A. L.; Macleod, A.; Noppen, S.; Sanderson, J.; Guérin. C. J. Colocalization Analysis in Fluorescence Micrographs: Verification of a More Accurate Calculation of Pearson's Correlation Coefficient. Microsc Microanal. 2010, 16, 710-724. 27


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

52. Di, H.; Liu, H.; Li, M.; Li, J.; Liu, D. High-Precision Profiling of Sialic Acid Expression in Cancer Cells and Tissues Using Background-Free Surface-Enhanced Raman Scattering Tags. Anal. Chem. 2017, 89, 5874−5881. 53. Phillips, J. C.; Braun, R.; Wang, W.; Gumbart, J.; Tajkhorshid, E.; Villa, E.; Chipot, C.; Skeel, R. D.; Kalé, L.; Schulten, K. Scalable Molecular Dynamics with NAMD. J. Comput. Chem. 2005, 26, 1781−1802. 54. Vanommeslaeghe, K.; Hatcher, E.; Acharya, C.; Kundu, S.; Zhong, S.; Shim, J.; Darian, E.; Guvench, O.; Lopes, P.; Vorobyov, I.; Mackerell, A. D. CHARMM General Force Field: a Force Field for Drug-Like Molecules Compatible with the CHARMM All-Atom Additive Biological Force Fields. J. Comput. Chem. 2010, 31, 671−690. 55. Jorgensen, W. L.; Chandrasekhar, J.; Madura, J. D.; Impey, R. W.; Klein, M. L. Comparison of Simple Potential Functions for Simulating Liquid Water. J. Chem. Phys. 1983, 79, 926−935. 56. Uhlenbeck, G. E.,; Ornstein, L. S. On the Theory of the Brownian Motion. Phys. Rev. 1930, 36, 823−841. 57. Feller, S. E.; Zhang, Y.; Pastor, R. W.; Brooks, B. R. Constant Pressure Molecular Dynamics Simulation: the Langevin Piston Method. J. Chem. Phys. 1995, 103, 4613−4621. 58. Ryckaert, J.-P.; Ciccotti, G.; Berendsen, H. J. C. Numerical Integration of the Cartesian Equations of Motion of a System with Constraints: Molecular Dynamics of n-Alkanes. J. Comput. Phys. 1977, 23, 327−341. 59. Andersen, H. C. Rattle: a Velocity Version of the Shake Algorithm for Molecular Dynamics Calculations. J. Comput. Phys. 1983, 52, 24−34. 60. Miyamoto, S.; Kollman, P. A. Settle: An Analytical Version of the SHAKE and RATTLE Algorithm for Rigid Water Models. J. Comput. Chem. 1992, 13, 952−962. 61. Tuckerman, M.; Berne, B. J.; Martyna, G. J. Reversible Multiple Time Scale Molecular Dynamics. J. Chem. Phys. 1992, 97, 1990−2001. 62. Darden, T.; York, D.; Pedersen, L. Particle Mesh Ewald: An N•log(N) Method for Ewald Sums in Large Systems. J. Chem. Phys. 1993, 98, 10089−10092. 63. Humphrey, W.; Dalke, A.; Schulten, K. VMD: Visual Molecular Dynamics. J. Mol. Graph. Model. 1996, 14, 33−38. 64. Fu, H.; Zhang, H.; Chen, H.; Shao, X.; Chipot, C.; Cai, W. Zooming across the Free-Energy 28


Page 28 of 30

Page 29 of 30 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Nano

Landscape: Shaving Barriers, and Flooding Valleys. J. Phys. Chem. Lett. 2018, 9, 4738–4745.

29


ACS Nano 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

For TOC only

30


Page 30 of 30

Reversible Self-Assembly of Nanoprobes in Live Cells for Dynamic

Recommend Documents