Aptamer-Anchored Rubrene-Loaded Organic Nanoprobes for Cancer Cell Targeting and Imaging

Abstract

Open AccessCCS ChemistryCOMMUNICATION1 Aug 2019Aptamer-Anchored Rubrene-Loaded Organic Nanoprobes for Cancer Cell Targeting and Imaging Yuanyuan Li†, Chuansheng Li†, Tao Jiang, Lihua Zhou, Pengfei Zhang, Ryan T. K. Kwok, Jacky Wing Yip Lam, Ping Gong, Lintao Cai and Ben Zhong Tang Yuanyuan Li† Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) HKUST Shenzhen Research Institute, Shenzhen 518057 (China) †Y. Li and C. Li contributed equally to this work.Google Scholar More articles by this author , Chuansheng Li† Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) †Y. Li and C. Li contributed equally to this work.Google Scholar More articles by this author , Tao Jiang Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) Department of Pharmaceutical Sciences, Nanfang Hospital, Southern Medical University, Guangzhou 510515 (China) Google Scholar More articles by this author , Lihua Zhou Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) Google Scholar More articles by this author , Pengfei Zhang *Corresponding authors: E-mail Address: [email protected]; E-mail Address: [email protected] Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) Google Scholar More articles by this author , Ryan T. K. Kwok Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) HKUST Shenzhen Research Institute, Shenzhen 518057 (China) Google Scholar More articles by this author , Jacky Wing Yip Lam Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) HKUST Shenzhen Research Institute, Shenzhen 518057 (China) Google Scholar More articles by this author , Ping Gong Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) Department of Pharmaceutical Sciences, Nanfang Hospital, Southern Medical University, Guangzhou 510515 (China) Google Scholar More articles by this author , Lintao Cai Guangdong Key Laboratory of Nanomedicine, CAS Key Laboratory of Health Informatics, Shenzhen Bioactive Materials Engineering Lab for Medicine, Institute of Biomedicine and Biotechnology. Shenzhen Institutes of Advanced Technology, Chinese Academy of Sciences, Shenzhen 518055 (China) Google Scholar More articles by this author and Ben Zhong Tang *Corresponding authors: E-mail Address: [email protected]; E-mail Address: [email protected] Department of Chemistry, The Hong Kong Branch of Chinese National Engineering Research Center for Tissue Restoration and Reconstruction, Institute for Advanced Study, Department of Chemical and Biological Engineering, The Hong Kong University of Science and Technology, Clear Water Bay, Kowloon (China) HKUST Shenzhen Research Institute, Shenzhen 518057 (China) Center for Aggregation-Induced Emission, NSFC Center for Luminescence from Molecular Aggregates, SCUT-HKUST Joint Research Laboratory, State Key Laboratory of Luminescent Materials and Devices, South China University of Technology, Guangzhou 510640 (China) Google Scholar More articles by this author https://doi.org/10.31635/ccschem.019.20180037 SectionsSupplemental MaterialAboutAbstractPDF ToolsAdd to favoritesDownload CitationsTrack Citations ShareFacebookTwitterLinked InEmail Fluorescence imaging has become an indispensable technique in cancer research because it can reveal informative molecular, cellular, anatomical, and functional insights. Development of advanced fluorescent probes with superior sensitivity and biological selectivity for fluorescence imaging is thus imperative. To move forward in this direction, we developed an easy self-assembly method for fabricating aptamer-anchored rubrene-loaded organic fluorescent nanoprobes. The aptamer-modified organic nanoprobes integrated the best features of the organic light-emitting materials and the aptamers, thus endowing them with excellent cell-targeting capability, high stability, and good biocompatibility. By using this general method, a variety of biocompatible and highly bright organic fluorescent nanoprobes based on novel organic light-emitting materials with specific recognition could be easily constructed for real-time biosensing and long-term biomedical imaging. Download figure Download PowerPoint Introduction Cancer has become one of the biggest burdens of disease to humans. Implementation of early detection and development of targeted therapies are essential to reduce the high mortality rates associated with cancers. To achieve these goals, molecular probes capable of recognizing cancer cell-specific targets are needed.1 The use of recognition motifs, such as folic acid, peptide, and antibodies for specific tumor accumulation and active binding, is considered one of the most promising approaches for cancer targeting.2 Because of the high affinity, specificity, and wide range of target availability, monoclonal antibodies (mAbs) have been the workhorse of molecular targeting.3 However, production of the mAbs is costly and time consuming, and their manipulation requires particular controls and conditions because of easy protein denaturation, which may cause specific activity loss. Alternatively, aptamers are single-stranded DNA or RNA oligonucleotides that can bind to their target molecules (such as proteins, drugs, small molecules, and cells) with high affinity and selectivity by folding into distinct secondary and tertiary structures.4 Compared with mAbs, aptamers demonstrate several advantages, such as their relatively small size, lack of immunogenicity, and the ease of synthesis and modification.5 All these merits make aptamers promising recognition motifs to modify nanoparticles and lead to their increasing use for specific cancer imaging and therapy.6–9 Fluorescence imaging has become an indispensable technique in cancer research because it can reveal informative molecular, cellular, anatomical, and functional insights.10,11 Because of high sensitivity and high visibility, fluorescence imaging systems are particularly suited for early detection of cancers, assessing tumor margins, and monitoring response to therapy.12,13 Despite the rapid advances in fluorescence materials and new imaging technologies, fluorescent probes with better performance, especially in sensitivity and biological selectivity, are highly pursued.14 Since the development of nanotechnology expanded rapidly in the past few decades, various nanomaterials with excellent optical properties have been synthesized and developed as novel nanoprobes (NPs) for fluorescence imaging.15 Compared with traditional organic small-molecule dyes, fluorescent nanoparticles exhibit brighter fluorescence, higher photostability, and larger absorption coefficients, and they provide a scaffold for attaching multiple ligand molecules and for targeting and imaging.16 However, most conventional fluorescent nanoparticles, such as semiconductor quantum dots, are composed of inorganic species, and even some toxic heavy metal cations, though their long-term toxicity remains a concern, which has greatly limited the further clinical transition of such types of NPs.17,18 In terms of biocompatibility, organic nanoparticles consisting of organic dye molecules or luminescent polymers encapsulated inside or decorated on the matrix of biocompatible polymers are generally considered to be less toxic than inorganic nanoparticles.19–22 Along with the rapid development of organic optoelectronics, such as organic field-effect transistors, organic solar cells, and organic light-emitting diodes, a number of high-performance organic light-emitting materials have been designed and studied.23,24 However, most of these kinds of materials were processed in organic solvents and showed obvious hydrophobic properties. To solve this thorny problem, in this work, we aim to develop a simple method to fabricate aptamer-targeting organic fluorescence NPs based on self-assembling strategies. By integrating the advantages of organic light-emitting material-based nanoparticles with the cell-targeting capabilities of aptamers, aptamer-functionalized organic NPs may open a new path to sophisticated design solutions for biomedical applications. Rubrene, a highly emissive organic semiconductor molecule, was chosen for fabrication of organic nanoparticles. As shown in Scheme 1, the aptamer–rubrene nanoprobes (Apt–Rub NPs) were self-assembled from a mixture of rubrene, 1,2-Distearoyl-sn-glycerol-3-phosphoethanolamine-N-maleimide (polyethylene glycol 2000) (PEG-lipid), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (lipid), and aptamer-cholesterol through a single-step and one-pot sonication method.25 The hydrophobic lipid segments tend to be embedded in hydrophobic rubrene aggregates, whereas the hydrophilic PEG chains extend into the aqueous phase to render the nanodots with water dispensability and enhanced colloidal stability. Starting from this scaffold and incorporating DNA aptamers to incorporate targeting functionality could potentially facilitate the translation of organic light-emitting materials to clinical practice. The cholesterol covalently linked to an aptamer served as an anchor to be firmly inserted into the lipid layer via hydrophobic interactions, and it enhanced the stability of the lipid layers. Cancer cell-specific targeting and imaging were successfully demonstrated using AS1411 aptamer-modified NPs, a 26-mer single-strand DNA that has been confirmed to selectively bind to nucleolin (which is overexpressed on the membranes of several cancer cells but is absent in normal cells).26 This kind of particularly high affinity between aptamer AS1411 and nucleolin could be used for improving the endocytosis of rubrene into cancer cells. Scheme 1 | Schematic illustration of preparation of the aptamer–rubrene nanoprobes (Apt–Rub NPs) by one-pot self-assembly method. Download figure Download PowerPoint Results and Discussion The Apt–Rub NPs were one-pot prepared by simply mixing the hydrophobic rubrene and lipid with the cholesterol-tagged aptamer during the hydration step of the lipid-nanoparticles formulation. The absorption and photoluminescence (PL) properties of rubrene and rubrene-loaded nanoparticles with or without aptamer anchoring dispersed in 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer were first investigated. As shown in Figure 1a, the free rubrene molecules in chloroform exhibited two absorption peaks at ∼480 and 530 nm. Identically, the absorption peaks for the rubrene nanoparticles with or without aptamer anchoring also exhibited two absorption peaks at 480 and 530 nm, indicating that the aptamer had no effect on the absorption profile of rubrene. The maximum emission of rubrene nanoparticles appeared at 560 nm, with an intense emission tail extending to 650 nm (Figure 1b), which held advantages of low optical absorption and autofluorescence from biomolecules. It is noteworthy that the emission spectra of the aptamer-anchored rubrene nanoparticles were similar to those of nanoparticles without aptamer functionalization. These data supported the idea that the rubrene maintained its optical properties after encapsulation with lipid and aptamer-cholesteryl. Toward the goal of achieving highly bright fluorescent NPs, it was critical to optimize the loading capability of rubrene to give the highest fluorescent signal. We first fabricated different benches of nanoparticles by changing the mass of free rubrene ranging from 37.5 to 100 μg. As the results show in , it indicated that the fluorescent intensity of the nanoparticles initially increased along with the increase of the rubrene loading. However, when the loading amount of rubrene was larger than 75 μg, the fluorescent intensity of the nanoparticles dropped, which was possibly due to the aggregation-caused quenching effect of rubrene. Therefore, we chose 75 μg as the optimal amount for the rest of the nanoparticle preparation in this work, and the loading efficiency of that was 3.7% (wt). The nanoparticles were further characterized using transmission electron microscopy (TEM) and dynamic light scattering (DLS) (Figure 1c and d). As the DLS data show in Figure 1c, the hydrodynamic size of rubrene-loaded nanoparticles with and without aptamer modification revealed an average hydrodynamic diameter of ∼73 and ∼92 nm, respectively. The diameter of as-prepared Apt–Rub NPs from DLS analysis was about 20 nm larger than that of rubrene NPs, suggesting that the attachment of aptamers increased the hydration diameter of the NPs. The TEM results, however, revealed that the as-prepared Apt–Rub NPs were in a spherical shape with an average size of ∼40 nm. The average size estimated by DLS was larger than that measured by TEM because the NPs had a certain extent of swelling and hydration in solution.27 The surfaces of rubrene NPs were negatively charged (−22.4 mV), which was ascribed to the negatively charged phosphate group on the hydrophilic head of phospholipid. After modification with aptamers, the zeta potential of the Apt–Rub NPs further decreased to about −26.6 mV(). To further confirm that the aptamers have successfully anchored on the NPs, a complementary DNA (cDNA) to the aptamer sequence immobilized on a microbead was prepared through streptavidin–biotin interactions (Figure 2a and ). Then, the NPs with or without aptamer anchors were mixed with the cDNA-decorated microbeads and imaged with confocal microscopy. As illustrated in Figure 2b, the Apt–Rub NPs showed bright emission on the surface of microbeads. On the contrary, there was an undetectable fluorescent signal on the microbeads for the rubrene NPs and the blank ones. Figure 1 | Characterization of rubrene NPs and Apt–Rub NPs. (a and b) Normalized absorption spectra and normalized PL spectra of free rubrene molecules dissolved in chloroform and rubrene NPs and Apt–Rub NPs dispersed in deionized water. Excitation wavelength: 488 nm. (c and d) DLS measurements of the rubrene NPs and Apt–Rub NPs. Inset: TEM images of the rubrene NPs and Apt–Rub NPs. Scale bar: 100 nm. Download figure Download PowerPoint Figure 2 | (a) Schematic illustration of the hybridization of aptamer-anchored rubrene NPs with its cDNA immobilized on a microbead. (b) Confocal imaging of the hybridization of Apt–Rub NPs with the cDNA immobilized on the microbead. Excitation wavelength: 488 nm; emission wavelength: 580–700 nm. Scale bar: 100 μm. Download figure Download PowerPoint Because the conditions in cell or tumor microenvironments can be significantly different, it is critical to evaluate the fluorescence intensity fluctuation of the Apt–Rub NPs under various conditions before proceeding to cellular studies. As depicted, the fluorescence (Figure 3) and hydrodynamic diameter () stability of as-prepared NPs was evaluated under several biologically relevant conditions, including (1) the pH range 3–9, (2) in different ionic strength buffers (0, 0.5, 1, and 2 M NaCl), (3) in the presence of biomolecules and culture media, and (4) in the presence of various ions under room temperature with light exposure conditions. The results showed that all the NP samples stayed stable with no sign of aggregation or loss of fluorescence. Evidently, the as-prepared Apt–Rub NPs were highly stable. Figure 3 | Stability of Apt–Rub NPs in different conditions. PL intensities of the Apt–Rub NPs at 630 nm (a) in buffer solutions at pH from 3 to 9, (b) in the neutral buffer solution with different concentrations of NaCl, (c) in cell culture medium, and (d) in the presence of various ions. Inset: Fluorescence images of the corresponding Apt–Rub NPs in different conditions taken under 365 nm UV light illumination. Download figure Download PowerPoint To demonstrate the targeting capability of Apt–Rub NPs, AS1411 aptamer, a 26-base guanine-rich DNA aptamer, was chosen as the model targeted cell recognition ligand embedded in the self-assembled organic dots. The AS1411 aptamer has been discovered to have high binding affinity to nucleolin, a nucleolar phosphoprotein overexpressed on the surface of several cancer cell lines (such as breast cancer cells and lung cancer cells).28 To determine the specificity of as-prepared Apt–Rub NPs for targeting nucleolin, cancer cell lines such as MCF-7 (human breast cancer cell line), C6 (rat glial tumor cell line), and A549 cells (human lung cancer cell line), and the normal healthy cells, 293T cells (human embryonic kidney cell line) were incubated with Apt–Rub NPs and rubrene NPs, separately. The fluorescence of the treated cells was collected using laser confocal fluorescence microscopy and flow cytometry (Figure 4). As illustrated in Figure 4a, MCF-7, C6, and A549 cells treated with Apt–Rub NPs were highly fluorescent when compared with those treated with the rubrene NP controls. By contrast, for the normal cell line 293T, no fluorescence intensity was detected treated either with Apt–Rub NPs or rubrene NPs. The results demonstrated that the Apt–Rub NPs can selectively bind to the nucleolin-overexpressed cancer cells (MCF-7, C6, and A549 cells) more than to the normal cells (293T). As a control, the random DNA-modified rubrene NPs were incubated with A549 cells (), and the results showed that there was a little signal on cancer cells, which further confirmed the targeting ability of the aptamer on nanoparticles. To avoid the nonspecific binding of the NPs, we have measured the cellular uptake at 0 °C. The confocal imaging results showed in that there was negligible uptake because of the energy-dependent uptake by the NPs through endocytosis pathways.29,30 In addition, the fluorescence of the treated cells was further analyzed by the flow cytometry (Figure 4b). The Apt–Rub NPs were specifically internalized by cells only with overexpression of nucleolin on the cell membrane (MCF-7, C6, and A549 cells). The flow cytometric analysis results were consistent with previous fluorescent imaging, collectively suggesting that the AS1411-modified Apt–Rub NPs can target cancer cells selectively and efficiently. The bioactive AS1411 aptamer endowed the Apt–Rub NPs, with the capability to specifically and exactly recognize nucleolin on the cancer cells. Apart from the imaging performance, biocompatibility is the main concern for the imaging contrast during its use in biomedical applications. The cytotoxicity of the Apt–Rub NPs after incubation of 24 h with several cell lines (MCF-7, C6, A549, and 293T cells) was evaluated using a Cell Counting Kit-8 (CCK-8) assay (Figure 5). The results demonstrated that the as-prepared Apt–Rub NPs had no obvious cytotoxicity toward all the cell lines at different concentrations. To evaluate the preliminary long-term toxicity of rubrene NP, we monitored the weight fluctuation and behaviors of the mouse after injection (). There were no abnormalities in activity, drinking, and eating. At the same time, the body weight of the treated group increased gradually in a manner similar to that of the control group. Figure 4 | Specific cancer cell-targeting experiments of the Apt–Rub NPs. MCF-7, C6, A549, and 293T cells were cultured with Apt–Rub NPs (concentration: 100 μg/mL) at 37 °C for 60 min, followed by washing with PBS. Rubrene NPs and blanks were used as controls. (a) Confocal microscopy imaging and (b) flow cytometric analysis of the cellular uptake of Apt–Rub NPs, rubrene NPs, and blank. Scale bar: 50 μm. Objective: 63×; excitation wavelength: 488 nm; emission wavelength: 580–700 nm. Download figure Download PowerPoint Figure 5 | Cell viability of the Apt–Rub NPs in MCF-7, C6, A549, and 293T cell lines at various concentrations ranging from 62.5 to 1,000 μg/mL after 24 h of incubation. Data represent the mean value ± standard deviation, n = 5. Download figure Download PowerPoint Conclusion In summary, we developed a simple one-pot method to fabricate aptamer-modified rubrene-loaded nanoprobes (Apt–Rub NPs) for targeted cancer imaging. This one-step method is based on self-assembly and hydrophobic interactions between the cholesterol and the lipids. As-prepared Apt–Rub NPs showed specific targeting capability, high stability, and good biocompatibility. Through this universal approach, various aptamer-modified organic photoelectric material-based NPs with different sequences and functions could be designed and fabricated. The diversity of aptamers and the huge amount of organic photoelectric materials provide unlimited possibilities to construct biocompatible organic fluorescent NPs with specific recognition and highly sensitive tracking, which can be further used for high throughput, rapid and reproducible biosensing detection, and biomedical imaging. Compared with antibodies, aptamers have higher resistance to deactivation. Therefore, aptamer-decorated organic NPs are suitable for long-term, real time, and dynamic sensing, tracking, and imaging, making them extremely promising in biomedical and bioanalytical applications. Experimental Sections Materials 1,2-Distearoyl-sn-glycerol-3-phosphoethanolamine-N-maleimide (polyethylene glycol 2000) (DSPE-PEG) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) were obtained from Avanti Polar Lipids (United States). Hoechst 33342, Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), Dulbecco’s phosphate buffered saline (PBS), trypsin EDTA (0.5% trypsin, 5.3 mM EDTA tetrasodium), and penicillin–streptomycin (100 U/mL) were purchased from Life Technologies. Ultrapure water was obtained from a Millipore-Q system. The sequence of aptamer-cholesterol used in this study was 5′-GGT GGT GGT GGT TGT GGT GGT GGT GGT TTT TTT TTT TT-cholesterol-3′ and synthesized by Sangon Biotech Co. Ltd (Shanghai, China). As control, random DNA sequence 5′-GCACTGGTCGGCCATGGGTAGCGACGGTCCCTAACGTT-cholesterol-3′ was synthesized. CCK-8, the cell proliferation cytotoxicity assay kit, was obtained from Fanbo Biochemicals. Unless otherwise noted, all chemicals were obtained from Sigma-Aldrich and were used as received. Instruments UV-Visible (UV-vis) absorption spectra were taken on a PerkinElmer Lambda 25 UV/Vis absorption spectrophotometer. PL spectra were recorded with an Edinburgh F900 fluorescent spectrometer. TEM images were taken on JEM 100CXII (JEOL). The TEM samples were prepared by placing a drop of Apt–Rub NP solution onto a 300-mesh copper grid and then drying the sample at room temperature overnight. The particle sizes and zeta potential of particles were characterized on a Nano-Zetasizer (Nano ZS, Malvern; Malvern Instruments, United States) at 25 °C. Fabrication and characterization of Apt–Rub NPs Stock solutions of rubrene with different amounts (37.5–100 µg) of DPPC (1.25 mg), cholesterol (0.325 mg), and DSPE-PEG (0.38 mg) in chloroform were mixed in a scintillation vial. The mixture was blow-dried with N2 and further dried under vacuum overnight. After complete evaporation of the chloroform, the residue was heated at 50 °C. The preparation buffer (5 mL), containing 25 mM 2-[4-(2-hydroxyethyl)-1-piperazinyl]-ethane sulfonic acid (HEPES, pH 7.4), 150 mM NaCl, 5 mM KCl, 1 mM MgCl2, and 1 mM CaCl2 was added to the dried lipids followed by addition of 100 nmol of cholesterol-tagged DNA aptamer. The mixture was incubated for 6 h at 50 °C and further incubated for 10 min at 37 °C during the sonication. The desired Apt–Rub NP was stocked at 4 °C. In vitro stability tests Fluorescent stability assays were carried out at different pH levels with different ions and buffer mediums. Ten microliters of Apt–Rub NPs were separately added in 490 µL PBS solution with pH levels ranging from 3 to 9; fluorescent intensity was measured to evaluate its stability to pH. Other stability tests were also carried out via the same method while changing the diluents to high concentrations of NaCl (0, 0.5, 1, and 2 M) or 0.1 M different kinds of metal ions (Na+, K+, Mg2+, Ca2+, Cd2+, Cu2+, Mn2+, Co2+, Ni2+, Zn2+, Fe3+, Cl−, NO3−, CO32−, SO42−, and PO43−). Hydrodynamic size assays were carried out at different pH levels with different ions and buffer mediums. Ten microliters of Apt–Rub NPs were separately added in a 490 µL PBS solution with pH levels ranging from 3 to 9. Hydrodynamic size was measured to evaluate its stability to pH. Other stability tests were also carried out via the same method while changing the diluents to high concentrations of NaCl (0, 0.5, 1, and 2 M) or 0.1 M different kinds of metal ions (Na+, K+, Mg2+, Ca2+, Cd2+, Cu2+, Mn2+, Co2+, Ni2+, Zn2+, Fe3+, Cl−, NO3−, CO32−, SO42−, and PO43−). Cell culture MCF-7, C6, A549, and 293T cells were purchased from ATCC. The four cell lines were cultured in DMEM with 1% penicillin–streptomycin and 10% FBS, at 37 °C in a humidified incubator with 5% CO2. The culture medium was changed every 2 days, and then the cells were treated with 0.25% trypsin EDTA solution after reaching confluence. Cellular uptake analysis MCF-7, C6, A549, and 293T cells were seeded into eight-well chambered cover glasses (Lab-Tek, Nunc, United States) in 200 µL of medium. After 24 h of culturing at 37 °C under the condition of 5% CO2, the medium was replaced by a new medium with as-prepared Apt–Rub NPs or rubrene NPs. After 2 h of incubation, the cells were washed three times and maintained in HEPES buffer. Finally, the cells were imaged by confocal laser scanning microscope (TCS SP5, Leica, Germany) equipped with argon, red HeNe, and green HeNe lasers. Images were collected using a Plan-apochromat 63×/1.4 oil immersion objective by sequential line scanning, with excitation at 405 and 488 nm, along with a bright-field image. Emission was collected by photomultiplier tubes in the ranges 423–492 and 510–570 nm, obtained by tunable high-reflectance mirrors. The cell uptake was further quantitatively evaluated by flow cytometry. MCF-7, C6, A549, and 293T cells were seeded into six-well plates (105 cells per well) in 1 mL medium. After 24 h, the original medium was removed and new medium containing Apt–Rub NPs was added into each well. A medium with rubrene NPs was used as a negative control. After 1 h of incubation, the cells were harvested by 0.05% trypsin EDTA. The emission was detected by flow cytometer (Accuri C6, BD, United States). Cytotoxicity assay CCK-8-based cell viability assay was performed to assess the metabolic activity of cells. Cytotoxicity assay was acquired on a multifunctional microplate reader (BioTek Inc., Winooski, VT). Cells were seeded in 96-well plates (Costar, IL, United States) at an intensity of 6 × 104 cells/mL. After 24 h of incubation, the medium was replaced by the rubrene NPs/Apt–Rub NPs in media at different concentrations, and the cells were then incubated for 24 h. After the designated time intervals, 10 μL of CCK-8 reagent was added into each well. After 3 h of incubation at 37 °C, the absorbance of CCK-8 at 450 nm was monitored by the microplate reader. Cell viability was expressed by the ratio of absorbance of the cells incubated with nanoparticle solution to that of the cells incubated with culture medium only. Supporting Information Supporting information is available. Acknowledgments This work was partially supported by the University Grants Committee of Hong Kong (AoE/P-03/08), the Research Grants Council of Hong Kong (16301614, 16305015, and N_HKUST604/14), and the Innovation and Technology Commission (ITC-CNERC14SC01, ITS/254117, and RE:ITCPD/17-9). B. Z. T. is grateful for the support from the Guangdong Innovative Research Team Program of China (201101C0105067115). P. F. Z. is grateful for the support from the National Natural Science Foundation of China (Grant No. 81501591).