Novel Enhancer for Luminol-AuNP Electrochemiluminescence and

Oct 19, 2018 - The detection limit of the target cell amounts reached level 50. The specificity test and recovery test showed that the method had a go...
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Novel Enhancer for Luminol-AuNP Electrochemiluminescence and Decoration on RNA Membranes for Effective Cytosensing Yingshu Guo,*,†,‡,§,¶ Xinxin Shang,†,‡ Fei Liu,†,¶ Yinhua Hu,†,‡ Shuang Li,†,‡ Jia Liu,†,§ and Fei Wu†

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Shandong Province Key Laboratory of Detection Technology for Tumor Makers, School of Chemistry and Chemical Engineering, Linyi University, Linyi 276005, China ‡ Shandong Provincial Key Laboratory of Life-Organic Analysis, College of Chemistry and Chemical Engineering, Qufu Normal University, Qufu 273165, China § Key Laboratory of Sensor Analysis of Tumor Marker, Ministry of Education, College of Chemistry and Molecular Engineering, Qingdao University of Science and Technology, Qingdao 266042, People’s Republic of China ¶ College of Chemistry, Chemical Engineering and Materials Science, Collaborative Innovation Center of Functionalized Probes for Chemical Imaging in Universities of Shandong, Key Laboratory of Molecular and Nano Probes, Ministry of Education, Institute of Biomedical Sciences, Shandong Normal University, Jinan 250014, People’s Republic of China S Supporting Information *

ABSTRACT: Herein, a simple and nice method to synthesize RNA membrane nanostructures for a controlled capture and cytosensing by aptamer and luminol-AuNPs (luAu NPs) is presented. The RNA membrane made up of many RNA transcripts was prepared by a one-pot reaction, the rolling circle transcription (RCT) process using rNTP and RNA polymerase, so the efficient RNA membrane could be obtained successfully at a much lower cost than the system used for RNA molecule synthesis commercially. The modifications of the NH2 group could not degrade the stability of RNA membrane in the serum. The prepared RNA membrane nanostructures showed interesting capture efficiencies toward the Ramos cell due to the effect of aptamer−ligand interactions on the nanostructures. Additionally, based on the detection of the luAu NP electrochemiluminescence (ECL) system, a novel enhancing method was employed by 3aminopropyl-triethoxysilane (APS) for luAu NP ECL, which has never been reported, to improve the sensitivity. The ECL intensity was proportional to the target cell amounts, and the dynamic range was 50−5000 cells. The detection limit of the target cell amounts reached level 50. The specificity test and recovery test showed that the method had a good selectivity, stability, and repeatability. Therefore, this method will provide a favorable analysis for biosensing. KEYWORDS: RNA membrane, cell capture, electrochemiluminescence, enhancer

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INTRODUCTION Circulating tumor cells (CTCs) are cells that1 are derived from primary tumors or metastatic tumors and acquire the ability to detach from the basement membrane and invade tumor cells that enter the circulatory system through the tissue matrix body and form metastases in important distal organs. In addition to the genome and proteome, tumor cells are information-carrying freighters of interest regarding tumor growth. Tumor cells are the source of malignant tumor metastasis.2 Therefore, the detection of tumor cells is of potential value in the diagnosis and prognosis of tumor metastasis, drug development, individualized treatment, and exploration of the tumor metastasis mechanism and is expected to be a new cancer “biomarker” for efficacy assessment and individualized targeted therapy.3,4 However, the elucidation of the function of tumor cells in physiological fluids is still far © XXXX American Chemical Society

behind that of the genome and proteome because tools are lacking for reliably identifying tumor cells. Several methods, including aptamer-mediated capture,5−8 antisense-triggered release,9 and immunocapture microfluidic platform,10,11 have been used for cancer cell recognition, capture, separation, and imaging and are considered promising tools for counting tumor cells. These techniques can recognize and bind rare tumor cells because of the design of interaction between bionanomaterial conjugates and typical tumor cell surface biomarkers. Tumor cells may have surface biomarkers.12 The abundance of a single surface biomarker may wave following Received: August 29, 2018 Accepted: October 19, 2018 Published: October 19, 2018 A

DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX

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partially complement each other for two kinds of long singlestranded RNAs. So the two kinds of long single-stranded RNAs are partially complementary and then form RNA membrane. This method is much less expensive than industrially synthesized RNA molecules. In particular, with our surface decoration, we have successfully produced RNA membranes with luAu NPs. We selected Ramos as a tumor cell model because lymphoma, a primary node or other lymphoid tissue of malignant tumors that originates in hematopoietic malignancies, can occur at any age.26 Advances in space and time for high payloads and RNA membrane costs enabled us to investigate new approaches to capture and inhibit tumor cells in blood using aptamer-binding strategies. However, no quantitative or qualitative analysis is available for the interaction between Ramos and the specific conjugates, and on the basis of the conceptual advantage, this study can contribute important information to the field of luAu NP ECL intensity.

cell cycle changes, which cause the binding affinity between the biomarker and its recognition molecule to change.13 RNA, a single-stranded long molecule, needs to form a certain secondary structure or even tertiary structure by the base pairing principle to perform biological functions. Because of the progress of RNA biology and RNA nanotechnology, RNA has been thought as a powerful building material for the rapidly developing field of synthetic biology over the years. The concept of RNA nanotechnology was first proposed in 1998.14 It refers to the design, preparation, and application of nanoRNA architecture by using RNA as the main framework through bottom-up self-assembly. For example, compared with ribosomes, splicing bodies, and even complex DNA nanostructures, one of the advantages of RNA nanotechnology is that it can scientifically design structures comparable to natural RNA materials in size and complexity. Particles, targeting ligands, therapeutic moieties, and regulators based on RNA nanotechnology can consist of RNA nucleotides. In these years, all sorts of nanoparticles, depending on the self-assembly of various multimeric RNA units, have been designed multiple times and with multiple types of cycle structure modules, such as 2D and 3D polygons, arrays, and filaments.15−17 In all of these years, various factors have hindered the extensive use of RNA as a building material, such as susceptibility to the degradation of RNA enzymes, sensitivity to disaggregation after systemic injection, and toxicity, and adverse immune reactions. The above problems were largely solved: the modifications of 2′-fluoro (2′-F) or 2′-O-methyl (2′-OMe) on the •OH group of the ribose can maintain the chemical stability of RNA in serum;18 certain naturally occurring linker sequences are thermodynamically stable and allow the all RNA nanoparticles to remain in good condition at ultralow concentrations,19,20 and the immunogenicity of RNA nanoparticles relies on the sequence and shape, so RNA nanoparticles can be modulated to excite the result of inflammatory cytokines21 or make the RNA nanoparticles nonimmunogenic and nontoxic by intravenous injection of 30 mg/kg repeatedly.22 RNA nanotechnology will make exosome RNA play a vital role in the research and treatment of living cells and organisms.23,24 Lately, rolling circle transcription (RCT), an isothermal enzymatic nucleic acid amplification method, enables large-scale synthesis of RNA nanostructures. The principle of RCT is to convert dNTPs into single-stranded RNA under the action of enzymes by using a circular DNA as a template and a short DNA primer (complementary to a partial circle template), which contains hundreds of repeated template complementary fragments. A rapid isothermal reaction of RCT technology has a great application value and potential in cell imaging, drug delivery, nucleic acid detection, and so on. Recently, we reported folic acid-modified RNA nanoflowers for folate receptor-positive cell recognition. Our technique realized the surface modification of RNA nanoflowers to activate its surface, overcoming the characteristics of a poor selectivity.25 This research applies this new idea to achieve mass production of a large RNA membrane (Scheme S1) containing abundant amino acids. The RNA membrane made up of many RNA transcripts was prepared by a one-pot reaction, the rolling circle transcription (RCT) process. In the RCT process, singlestranded RNA with a periodic sequence was synthesized by incubating a DNA circular template and four kinds of rNTP with the help of the T7 RNA polymerase. We present two types of template sequences (DNA 1 and DNA 2) that

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EXPERIMENTAL SECTION

Materials and Reagents. We obtained 1-ethyl-3-[3-(dimethylamino) propyl]-carbodiimide (EDC), N-hydroxysuccinimide ester (NHS), Tris(2-carboxyethyl)phosphine hydrochloride, and 3-aminopropyl-triethoxysilane (APS) from Sigma-Aldrich; streptavidinmodified 96-well culture plate and HAuCl4 from Sangon Biotech (Shanghai) Co., Ltd., (China); T7 RNA polymerase and ribonucleotide (dATP, dGTP, dCTP, NH2-dUTP) from New England Biolabs; and T4 DNA ligase from Thermo Scientific. Luminol was purchased from Sigma. We obtained all synthesized oligonucleotides from Sangon Biotech Co., Ltd. (Shanghai, China) (Table S2). The experimental water was sterilized water without nuclease. Apparatus. We used a transmission electron microscope (TEM) (JEM-2100, JEOL) and scanning electron microscope (SEM) to characterize morphologies. We obtained the UV−vis absorption spectrum from a UV−vis spectrophotometer (Cary 60, Agilent), conducted cyclic voltammetric (CV) experiments using a CHI600E electrochemical workstation (Shanghai, China), and measured ECL intensities using a model MPI-E electrochemiluminescence analysis system (Xi’an Remex Electronics, Xi’an, China). Preparation of luAu NPs. In order to prepare luAu NPs, 50 mL of HAuCl4 solution (0.01%, w/w) was boiled and stirred constantly; then 0.8 mL of 0.01 mol/L luminol solution was added. At the boiling point, the color of the mixture changed from cream to purple or redpurple, and then the mixture cooled to room temperature and was characterized by TEM and UV−vis spectra. Cell Capture and Release. For this experiment, the 96-well plate was modified with streptavidin. First, we synthesized the circular DNA 1 by hybridizing DNA 1 and primer 3. We synthesized the circular DNA 2 by hybridizing DNA 2 and primer 2. We then seeded the circular DNA 1/primer 3 into the 96-well plate through the interaction between biotin and avidin for 12 h at 37 °C and added the circular DNA 2/primer 2, 4 mM of ribonucleotide solution mix (dATP, dGTP, dCTP, NH2-dUTP), and 5 U μL−1 of T7 RNA polymerase to react with the circular DNA 1 for 1 day at 37 °C for RNA membrane self-assembly. During this period, the RNA membrane-modified 96-well plate reacted with the luAu NP solution for 12 h at 4 °C and immobilized the luAu NPs onto the surface of the membrane via the Au−N bonds. We thoroughly rinsed the luAu NP/ RNA membrane-modified 96-well plate with ultrapure water to remove the free luAu NPs and then incubated the plate with 20 μM of Ramos aptamer solution in darkness for 12 h at 25 °C to enable the aptamer to self-assemble on the RNA membrane, after which we washed it to remove the free aptamer. We seeded Ramos (1 mL) with an initial density of 2 × 105 cells/mL into each well and washed the plate twice with PBS to remove unattached cells. To examine cell release, we incubated the 96-well plates in the solution containing 5 μM of secondary complementary sequences at 37 °C for 30 min and B

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Scheme 1. Aptamer- and luAu NP-Functionalized RNA Membrane for Cytosensing Based on APS as an ECL Enhancer

washed them twice to allow the released cells to fall off the membrane surface. Preparation of RNA Membrane-Modified Au Electrode. To pretreat the electrodes, we mirror polished the bare Au electrodes by 1.0, 0.3, and 0.05 mm of Al2O3 in turn and then processed them in ultrapure water and ethanol and again in ultrapure water for 10 min each. We then washed a Au electrode on a CHI600E electrochemical workstation at a scan rate of 100 mV s−1 with a cyclic voltammetry (CV) between −0.2 and 1.6 V (vs Ag/AgCl in 0.5 M H2SO4) until we obtained a reproducible cyclic voltammogram and characterized it by cycling between −0.2 and 0.6 V of Ag/AgCl in PBS (0.1 M, pH = 7.0) containing [Fe (CN)6]3 and [Fe (CN)6]4, obtaining a peak−peak difference of less than 100 mV. For the preparation experiment, after pretreating 5 μM primer 1 with 5 μL of 2 mM TCEP, we prepared the primer 1-modified Au electrode by fixing primer 1 on the Au electrode through the Au−S bonds for 12 h at 4 °C. We then cleaned the Au electrode slightly to remove unbound primer 1. We prepared the RNA membrane-modified Au electrode by immersing the circular DNA 1-modified Au electrode in the complementary circular DNA 2 solution mixed with 4 mM ribonucleotide solution mixture (dATP, dGTP, dCTP, NH2-dUTP) and 5 U μL−1 T7 RNA polymerase for 24 h at 37 °C. During this period, the circular DNA 1 and circular DNA 2 complemented each other to form the RNA membrane. We then washed the resulting RNA membrane-modified Au electrode thoroughly with ultrapure water to remove the free complementary circular DNA 2. Preparation of Aptamer/luAuNPs/RNA Membrane-Modified Au Electrode. During this period, we dipped the RNA membrane-modified Au electrode in luAu NP solution for 12 h at 4 °C and immobilized the luAu NPs onto the surface of the membrane with the Au−N bonds. We washed the luAu NP/RNA membranemodified Au electrode with ultrapure water to scour the free luAu NPs off, placed the modified electrode in luminol solution (1 × 10−3 mol L−1), and incubated it at 4 °C for 12 h, after which we immersed it in 20 μM Ramos aptamer solution in darkness for 12 h at 25 °C with the addition of 0.01 M EDC. After the aptamer self-assembled on the surface of the RNA membrane, we rinsed the Au electrode to remove the free aptamer. We then dripped 20 μL of deionized water, including 0.5 μL of APS, onto the surface of the Au electrode for 12 h at 25 °C.

primer 1 was attached to the gold electrode by the Au−S bond. Catalyzed by T4 DNA ligase, the 5′ phosphorylated DNA 1 molecule forms circular DNA with the help of primer 1. Also, the nick of DNA 2 was chemically closed based on the same principles of hybridization with one end of primer 2. The other end of primer 2 is designed to hybridization with primer 1. So, circular DNA 1 and circular DNA 2 are all captured on the gold electrode and used as the template for RCT in in vitro transcription with the help of the T7 RNA polymerase and ribonucleotide solution mix (dATP, dGTP, dCTP, NH2dUTP). The two kinds of RNA transcripts, which consisted of numerous contiguous periodic complements of DNA 1 or DNA 2, respectively, could be hybridized to form an aminoRNA membrane on a gold electrode. Then the luAu NPs and carboxyl-aptamer are immobilized onto the surface of the membrane with the Au−N bonds or amide bond. There is an ECL emission in the existence of coreactant, H2O2, resulting from the ECL reaction of luAu NPs. Ramos cells are captured by the aptamer, leading to the perturbation of ECL intensity. Accordingly, the as-proposed strategy can realize cytosensing and provides a universal platform for aptamer-related biomolecules. Cell Capture and Release. Surface-functionalized micro/ nanoscale particulates and particles are used extensively for separating cells and proteins. Enhanced local biomolecule− ligand interactions between cells and substrate can significantly improve capture efficiency. For specific cells, we coated the RNA membrane with aptamers using carboxyl aminoconjugated chemistry, enabling it to recognize and capture desired cells. We seeded Ramos (1 mL) with an initial density of 2 × 105 cells/mL into the aptamer/luAu NP/RNA membrane-modified 96-well plate (Scheme 2) and washed the plate twice with PBS to remove unattached cells. To measure cell capturing, we took the 10 μL of unattached cells, put it into the counting chamber of the counting board, and detected it with the LUNA-II automated cell counter (Logos Biosystems, Korea). Capture efficiency was 93%. Then, we seeded Ramos (1 mL) with an initial density of 2 × 105 cells/ mL in a 96-well plate and washed the plate twice with PBS to remove unattached cells. Experiments showed that a 96-well plate cannot capture cells (Figure 1B, inset). We also studied

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RESULTS AND DISCUSSION Principle. Scheme 1 illustrates the whole thinking. Partially complementary two linear single DNA strands (DNA 1 and DNA 2) were phosphorylated at the 5′-end. First, thiolated C

DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX

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aptamer/luAu NP/RNA membrane hardly captured HeLa cells. The same method was used to investigate the capture of the aptamer/luAu NP/RNA membrane for K562 cells and red blood cells, and we obtained similar results as the HeLa cells. The RNA membrane can capture Ramos cells efficiently, while other blood cells, including unwanted cells, freely cross the RNA membrane. The results furnish overwhelming evidence that aptamers and nanomaterials can be effectively bound. Both Ramos cells and HeLa cells demonstrated high survival rates (over 91%) (Figures S2) after culturing with the RNA membrane, indicating that the RNA membrane has an excellent biocompatibility. The drug doxorubicin (DOX), a chemotherapeutic, has the ability to bind in combination with double-stranded 5′-GC-3′ or 5′-CG-3′.27−29 The RNA membrane could provide many loading sites for DOX. After DOX was loaded in the functionalized RNA membrane, we observed the obvious inhibition of Ramos cell proliferation (46%) and slight inhibition of HeLa cell proliferation (84%) (Figure S3). This phenomenon occurs because the local high concentration of DOX in the RNA membrane can result in Ramos cells living in the DOX-enriched RNA membrane with a low cell viability. The aptamer attached to the RNA membrane could recognize Ramos cells specifically, and Ramos cells were captured, while HeLa cell were not. Therefore, the functionalized RNA membrane can clearly increase the conjugate specificity to recognize and capture specific tumor cells and further reduce the activity of the captured tumor cells through drug accumulations. To examine cell release, we incubated the 96-well plates in the solution containing 5 μM of the secondary complementary

Scheme 2. RNA Membrane for Cell Capture and Release

the capture ability of this RNA membrane to other cells. We seeded HeLa (100 μL) with an initial density of 1 × 104 cells/ mL into the aptamer/luAu NP/RNA membrane-modified 96well plate and washed the plate twice with PBS to remove unattached cells. To measure the cell capturing, we took the 10 μL of supernatant, put it into the counting chamber of the counting board, and detected it with the LUNA-II automated cell counter. The cell concentration of 10 μL of samples was 9.96 × 103 cells/mL. The experimental results showed that the

Figure 1. (A) Fluorescence images of cell capture. To ensure a clear observation, a Vybrant cell-labeling solution was used. (B) The number of cells after being captured and released on the RNA membrane. Inset: The number of cells after being captured and released with no RNA membrane. Error bars indicate standard deviations (n = 5). RSD was about 6%. D

DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX

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Figure 2. TEM images of the RNA membrane (A), luAu NPs (B), and luAu NP/RNA membrane (C). The UV−vis absorption spectra (D) of the RNA membrane (a), luAu NPs (b), and luAu NP/RNA membrane (c). Inset: a larger version.

Figure 3. (A) CV profiles of the (a) bare Au electrode; (b) DNA 1/primer 1 + Au electrode; (c) RNA membrane + Au electrode; (d) aptamer + luAu NPs + RNA membrane + Au electrode; and (e) cell + aptamer + luAu NPs + RNA membrane + Au electrode. (B) EIS of the (a) bare Au electrode; (b) DNA 1/primer 1 + Au electrode; (c) RNA membrane + Au electrode; (d) aptamer + luAu NPs + RNA membrane + Au electrode; and (e) cell + aptamer + luAu NPs + RNA membrane + Au electrode, in 10 mM PBS (2.5 mM Fe(CN)64‑/3− + 0.1 M KCl, pH 7.4).

sequences of DNA 3 at 37 °C for 30 min (antisense-triggered release) and washed it twice to allow the released cells to fall off the membrane surface. Photomicrographs showed that the Ramos cells adhered to the membrane until released after DNA 3 treatment (Figure 1A). The viability of the released cells was about 91%. Characterization of luAuNPs and RNA Membrane. We estimated the RCT reaction using polyacrylamide gel electrophoresis stained with ethidium bromide (EB), followed by

imaging under UV irradiation, as Figure S1 shows. Upon the addition of the ribonucleotide solution mixture and T7 RNA polymerase, the bright band of high molecular weight was observed in lane f, proving the RNA nanostructure product was fabricated. We used transmission electron microscopy (TEM) (JEM-2100, JEOL) to detect the morphologies of the RNA membranes, luAu NPs, and luAu NP/RNA membranes. As Figure 2 shows, the wrinkled and folded layer of the RNA membrane (Figure 2A) changed from the modification with E

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Figure 4. (A) The possible ECL mechanisms and ECL-potential curves of (a) Au NPs and (b) luAu NPs. (B) The possible ECL mechanisms and ECL-potential curves of (a) APS, (b) luAu NPs, and (c) luAu NP/APS-modified Au electrode in 0.1 M PBS (pH 8.0). Scan rate is 100 mVs−1.

luAu NPs (Figure 2B), and after further modification with ∼10 nm luAu NPs, we found that the interesting luAu NPs were densely decorated on the surface of the RNA membrane (Figure 2C). Figure 2D shows a comparison of the UV−vis spectra of the RNA membrane before and after modification with luAu NPs. The RNA membrane showed a peak at about 260 nm (Figure 2D, a), and luAu NPs showed peaks at about 355 and 525 nm (Figure 2D, b). After decoration with luAu NPs, we found new absorption bands at about 355 and 525 nm (Figure 2D, c), identified as the plasmon band of luAu NPs. This indicated the successful generation and attachment of luAu NPs on the RNA membrane-based substrate. The results agree with the TEM measurements. To investigate the stability of the NH2-modified RNA membrane (NH2-RNA membrane) in serum, we prepared the RNA membrane without the NH2-group modification (noNH2-RNA membrane) with the ribonucleotide solution mix (dATP, dGTP, dCTP, dUTP). After staining the two kinds of membranes with Sybr green II, we cultured all membranes in serum (5% or 10%). We monitored the degradation by measuring the fluorescent intensity of the Sybr green II released in solution to investigate the stability (Figure S4). The fluorescent intensity showed no significant increase during NH2-RNA membrane incubation in 5% serum and water after 3 h. The fluorescent intensity of two kinds of samples showed some increase when incubated in 10% serum and water after 6 h, suggesting that the 10% serum solution degraded the RNA membrane and NH2-RNA membrane after 6 h, suggesting that the NH2 could not degrade the stability of RNA membrane. Gel electrophoresis analysis (Figure S4C) showed that NH2RNA membrane was degraded partly in 10% serum solution after 6 h in accordance with fluorescent experiments (Figure S4B). Characterization of the Electrode. Cyclic voltammetry (CV) is an effective and facile electrochemical technique used mainly to determine the electrode surface microscopic reaction process. Figure 3A shows the cyclic voltammetry curves of the gradually improved Au electrode using 2.5 mM Fe(CN)64‑/3− as electroactive probes. Bare Au electrodes exhibit several intensive peaks and reversible redox peak currents in curve a, attributable to the enormous superficial area and good electrical conductivity. When DNA primer 1 was fixed on

the top of the electrode, there was a clear downward trend in the current signal (curve b). After the RNA membrane (curve c) formed, the peak current decreased gradually. However, when luAu NPs were deposited on the electrode modified by the RNA membranes, we observed a small increase in the ampere response (curve d). Because luAu NPs can increase the electrode’s effective superficial area, it was reasonable to increase the rate of electron transfer. The gap between the anodic and cathodic peaks widened after capturing cells via the specific binding between aptamer and cells (curve e). The appearance was attributable to the electron-inert character of RNA or DNA and cells, which impeded the electron transfer and mass transfer of Fe(CN)64‑/3− ions at the electrode surface. Electrochemical impedance spectroscopy (EIS) can get more impedance change on an Au electrode surface during the modification process. We performed EIS measurements on a CHI660 Electrochemical Analyzer (CH Instruments) in 0.01 M PBS (pH 7.4) containing 2.5 mM Fe(CN)64‑/3− using a three-electrode system: a modified Au electrode was used as the working electrode system, a platinum wire was used as the auxiliary electrode, and a Ag/AgCl electrode was used as the reference electrode. Curve a in Figure 3B shows the EIS of the bare Au electrode. A nearly straight line appeared, characterized by a mass diffusion limiting electron transfer process. With the immobilization of the DNA 1/primer 1 (b) and RNA membrane (c), the electric resistance of the electron transfer increased by degrees. However, when the aptamer and luAu NPs were fixed on the electrode, the EIS presented a lower resistance (d). The reason may be that luAu NPs anchored on the RNA membrane play an important role, similar to that of the wires, to facilitate electron transfer. One explanation for the mechanism of the Au−amines interaction assembly is due to the electrostatic adsorption of Au nanoparticles with a negative charge and protonated amines;30 the other reason is that it results from a strong covalent bond between amine and Au.31 In the last step, cell immobilization increased the resistance (e), confirming that the cells were captured. These phenomena are in accordance with the reality that the assembled tier prevents the electron transfer from the interface. Inference of the ECL Enhancement Mechanism. We can reasonably infer that the luminol is essential for the ECL production process from having observed no obvious ECL F

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Figure 5. (A) The ECL intensities of positive assay vs different Ramos cell numbers (a−i: 0, 50, 200, 1000, 2000, 3000, 4000, 4500, and 5000 cells) in 0.1 M PBS (pH 7.4). Inset: plots of positive ECL intensity vs Ramos cell numbers from 50 to 5000 cells. (B) Biosensor selectivity when analyzed with 1000 Ramos cells, 2000 K562 cells, 2000 HeLa cells, and 1000 Ramos cells in mixed solution. RSD was less than 5.4%.

former has an amine functional group, while the latter has an epoxy functional group, we can conclude that the amine group of APS plays an important role in ECL enhancement, which helps generate radicals during the ECL reaction. According to previous reports,34 APS’s reduction of species (RC3H7−NH2) on the electrode can produce oxidative species (RC3H7− NH2+•). In this case, O2 and APS redox on the electrode could produce O2−•, which can react with L−• and then further oxidize to AP*, causing subsequent light emission. Figure 4B describes the possible ECL mechanisms with their equations. Ramos Cell Detection. We assessed the cell biosensor’s sensitivity and quantitative range by culturing it with different kinds of cells (Scheme 1). Figure 5A indicates the relationship between the ECL response and various numbers of Ramos cells. The ECL signal decreased by piecemeal as the number of Ramos cells increased. We achieved a linear connection between the ECL intensity and the number of Ramos cells in a range from 50 to 5000 cells with a detection limit of 50 cells. We can show the linear relationship as Y = 12429.6−2.2X, with the correlation coefficient of R = 0.99, where Y is the ECL intensity and X is the number of Ramos cells. As shown in Figure 5B, we explored the biosensor’s selective recognition of Ramos cells by investigating 1000 Ramos cells, 2000 K562 cells, and 2000 HeLa cells. The results showed an ignorable cross-reactivity to Ramos, K562, HeLa cells, and the mixture in the presence of Ramos cells (1000 cells) with interfering cells despite the fact that they had high concentrations, showing that only matching cells could be captured and that the designed protocol’s selectivity to the target cells against other control cells was good. In order to investigate the repeatability of biosensors, the prepared five electrodes under the same conditions were used to analyze the cells with the same concentration, and the results showed a relative standard deviation (RSD) of no greater than 5.4%, indicating that the reproducibility of the functionalized RNA membrane ECL strategy was excellent. We have already confirmed the original Ramos cell aptamer sequence’s excellent specificity (see Figure 5B). In addition, we have verified that Ramos cells had a high specificity in pure buffer assays by studying their capacity of resisting disturbance with several other cells. In order to verify the specificity of the assay in 5% (v/v) whole blood, we examined nonspecific variations caused by various nontarget cells, performing all

signal when using Au NPs instead of luAu NPs (Figure 4A). The result revealed that the ECL came from the luminol and that the Au NPs could not generate ECL but rather increased the interface area to capture more luminol molecules and promote the electron transfer and the ECL production process. Additionally, the ECL emission intensity decreased when we used N2 to remove oxygen dissolved in the solution, indicating that dissolved oxygen was one of the important reactants for luAu NPs to produce ECL. Superoxide dismutase (SOD) is an important antioxidant enzyme in organisms. SOD has a special physiological activity and is the primary substance for scavenging free radicals in living organisms. It catalyzes the dismutation of the superoxide anion (O2−•) into hydrogen peroxide and molecular oxygen. Next, we added superoxide dismutase as a peroxide-species capture agent into an airsaturated detection solution, resulting in a remarkably decreased ECL intensity and indicating that O2−• was a crucial species participating in the ECL reaction. Oxidative species, such as O2 and O2−•, can enhance the ECL intensity. The results were concordant with the research demonstrating oxidizing substances of oxidized luminol anions to luminol radicals, such as O2−•.32,33 Previous works34 have potentially identified the mechanism as the oxidation of luminol anions − (LH ) from deprotonated luminol to luminol radicals (LH•), and the formation of LH• rapidly deprotonated to luminol monoanion radicals (L−•), with O2−• resulting from the reaction between L−• and dissolved O2. The luminol ECL production process originates in the exited state of 3aminophthalate (AP*), which the reaction of L−• with O2−• can induce. Our findings were consistent with those found in previous works. To identify the function of 3-aminopropyl-triethoxysilane (APS) for ECL enhancement, we performed control experiments. In the absence of luAu NPs, the ECL intensity of the APS in PBS was very weak (Figure 4B, curve a), which indicated that the APS could not produce ECL and, thus, the ECL was from the luAu NPs (Figure 4B, curve b). The result suggests that the APS only catalyzed the ECL reaction of luAu NPs. If the luAu NP-modified surface was coated with (3glycidyloxypropyl) triethoxysilane rather than APS, we observed no enhancement of ECL, which differs from the ECL enhancement of APS. Because APS and (3-glycidoxypropyl) triethoxysilane groups are similar, except that the G

DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX

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ACKNOWLEDGMENTS This work was supported by the National Natural Science Foundation of China (21575056), the Natural Science Foundation of Shandong Province of China (ZR2016JL010), and the Primary Research and Development Plan of Shandong Province (2018GSF118172).

experiments under the same conditions as those of the target analytes. This function is another advantage of this detection system. To study the applicability of the proposed system in biological fluids, we used human blood for spiking target cells with varied concentrations. The recoveries (between 93.3% and 110%) for the cells ranging from 1000 to 3000 were acceptable (Table S1). This study clearly demonstrates that the developed assay that was not compromised in blood could provide a potential analytical tool in real biological samples conditions. The results show that these nontarget cells have no significant effect on the ECL intensity even if the number of these cells is far higher than the number of the target analytes. This function is another advantage of this detection system.

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CONCLUSIONS We successfully developed an RNA membrane-based platform using oligonucleotides for cell-specific capture, release, and detection. The method is simple and has a high sensitivity and strong selectivity. This may be caused by the following four elements: (1) The RNA membrane was made up of many RNA transcripts and was prepared by a one-pot reaction, the rolling circle transcription (RCT) process using rNTP and RNA polymerase so that the efficient RNA membrane could be obtained successfully at a much lower cost than the system used for RNA molecules synthesized commercially. (2) An RNA membrane with the Ramos aptamer-toehold specifically recognized Ramos cells. (3) The whole process of intermolecular hybridization and transformation of the hybridized aptamer involves no factors that could destroy the cells. It can release living cells without damages for subsequent culture and live cell analysis. (4) We used APS, as a novel enhancer for luAu NP ECL, and discussed its enhancement mechanism. With the help of APS, the target cells can be detected in numbers as low as 50. Besides, this strategy could be easily extended to detect other cancer cells and biomarkers based on changing the Ramos aptamer to other ligands. Therefore, this programmable RNA-membrane platform has a huge potential for many biological and biomedical applications, such as diagnosis and prognosis of tumor metastasis, drug development, and individualized treatment. ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsabm.8b00478. Preparation of RNA membrane, recovery determined using the proposed cell assay, sequences used, experimental syntheses and methods, RCT reaction results, cell viabilities, and degradation of RNA membranes (PDF)

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Corresponding Author

*E-mail: [email protected]. Tel: +86-539-7258627. ORCID

Yingshu Guo: 0000-0002-6926-6152 Notes

The authors declare no competing financial interest. H

DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX

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DOI: 10.1021/acsabm.8b00478 ACS Appl. Bio Mater. XXXX, XXX, XXX−XXX