Design of a Modular DNA Triangular-Prism Sensor Enabling

Subscriber access provided by UB + Fachbibliothek Chemie | (FU-Bibliothekssystem)

Article

Design of Modular DNA Triangular-Prism Sensor Enabling Ratiometric and Multiplexed Biomolecule Detection on Single Microbead Yu Liu, Qiaoshu Chen, Jianbo Liu, Xiaohai Yang, Qiuping Guo, Li Li, Wei Liu, and Kemin Wang Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b04918 • Publication Date (Web): 28 Feb 2017 Downloaded from http://pubs.acs.org on March 1, 2017

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 free 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 accessible to all readers and 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.

Analytical Chemistry 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 8

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

Analytical Chemistry

Design of Modular DNA Triangular-Prism Sensor Enabling Ratiometric and Multiplexed Biomolecule Detection on Single Microbead Yu Liu, Qiaoshu Chen, Jianbo Liu*, Xiaohai Yang, Qiuping Guo, Li Li, Wei Liu, Kemin Wang* State Key Laboratory of Chemo/Biosensing and Chemometrics, College of Chemistry and Chemical Engineering, Key Laboratory for Bio-Nanotechnology and Molecular Engineering of Hunan Province, Hunan University, Changsha 410082, P. R. China. Tel.: 86-731-88821566. Fax: 86-731-88821566. E-mail: [email protected]; [email protected]. ABSTRACT: DNA nanostructures have emerged as powerful and versatile building blocks for the construction of programmable nanoscale structures and functional sensors for biomarker detection, disease diagnostics and therapy. Here we integrated multiple sensing modules into a single DNA 3D nanoarchitecture with a triangular-prism (TP) structure for ratiometric and multiplexed biomolecule detection on single microbead. In our design, the complementary hybridization of three clip sequences formed TP nanoassemblies in which the six single-strand regions in the top and bottom faces act as binding sites for different sensing modules, including an anchor module, reference sequence module and capture sequence module. The multifunctional modular TP nanostructures were thus exploited for ratiometric and multiplexed biomolecule detection on microbeads. Microbead imaging demonstrated that after ratiometric self-calibration analysis, the imaging deviations resulting from uneven fluorescent intensity distribution and differing probe concentrations were greatly reduced. The rigid nanostructure also conferred the TP as a framework for geometric positioning of different capture sequences. The inclusion of multiple targets led to the formation of sandwich hybridization structures that gave a readily detectable optical response at different ﬂuorescent channels and distinct fingerprint-like pattern arrays. This approach allowed us to discriminate multiplexed biomolecule targets in a simple and efficient fashion. In this module-designed strategy, the diversity of the controlled DNA assembly coupled with the geometrically well-defined rigid nanostructures of the TP assembly provides a flexible and reliable biosensing approach that shows great promise for biomedical applications.

The use of DNA nanostructures as nanoscale construction materials hold potential applications in the fields of molecular electronics,1,2 logic computation3,4 and diagnostic assays5 because of their easy programmability, sequence-based design, convenient chemical synthesis and facile integration.6,7 In particular, DNA nanostructures are considered to be one of the most important supports for the fabrication of functional nucleic acid sensors for disease biomarkers,8 biosensor development9,10 and biomedical imaging.11 Nucleic acid sensors often consist of several functional modules, including a capture module for the specific target recognition, a signal module for the sensing report, and an anchor module for the immobilization on the solid-phase interface.12 Based on their precise organization and intelligent nanoassembly, 3D DNA nanostructures can provide multiple binding sites, allowing for the strategic integration of multiple sensing modules. These geometrically well-defined rigid nanostructures also provide an excellent physical support and biocompatibility matrix that allows for spatial immobilization and oriented arrangement of the different modules.13 In recent years, a variety of multifunctional and controllable DNA nanostructures have been constructed using DNA nanotechnology for sensing applications.14-16 The sophisticated DNA origami has served as a molecular chip for the detection of single nucleotide polymorphisms, the study of the dynamic conformational switching of the G-Quadruplex, and as an addressable support for the precise positioning of proteins.17-20 A DNA tetrahedron and long-range self-assembled DNA

nanostructures have been developed for detection of microRNA, mRNA, and cocaine.21-24 In addition, DNA triangular-prism (TP) nanoarchitecture, with its geometrically welldefined rigid structure, can be intricately designed with several single-strand binding sites for nanoassembly. Sleiman et al. proposed the use of TP DNA cages for controlled assembly on spherically-supported lipid bilayers and used them to position functional components for transfer of molecular recognition information.25,26 Nie et al. also developed a reconfigurable 3D DNA TP nanostructure for the assembly of molecular logic circuits.27 Because of its well-defined 3D structure and convenient nanoassembly, DNA TP nanostructure is considered to be an excellent DNA framework candidate as a building block for the assembly of functional nanodevices. It is possible for us to make full use of the multiple binding sites of the DNA TP support to assemble different sensing modules together into a single device system for biomolecule detection. To date, several approaches have been developed for spatial orientation and positioning of different binding ligands with TP nanostructures.28 While assembly of different sensing modules on a DNA nanostructure might provide a novel basis for biosensing, we are unaware of any studies examining DNA nanostructures in combination with multiplex ratiometric assays.29,30 Here, a DNA TP nanostructure sensor was fabricated through a module integrating design strategy, which was exploited as a pattern sensing platform for ratiometric and multiplexed biomolecule detection. As shown in Figure 1, the

ACS Paragon Plus Environment


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 8

Figure 1. Integrating different sensing modules together into single TP nanostructure for ratiometric and multiplex assay. Hybridization of three clip sequences resulted in the formation of the TP (Step 1). The three capture sequences were fixed on the top face for target recognition (Step 2). A DEA-labelled sequence was trapped on the bottom face of the TP as a reference probe (Step 2). The anchor sequence was used for immobilization of the TP bracket on the microbeads (Step 3). Presence of the targets resulted in sandwich hybridization structure and the formation of TP5m on microbeads (Step 4). The strong fluorescent emission of the TP nanostructure on microbeads allowed for ratiometric analysis and multiplex biomolecule detection through pattern microbeads assay (Biosensing application).

box-like DNA TP was constructed with three 96-base clip sequences (C1, C2, C3). The two 10-base ends of the first clip (C1) hybridized to the back of the next clip (C2) through their complementary regions. Likewise, the second clip (C2) hybridized to the third clip (C3), and the third clip (C3) hybridized to the first clip (C1) to form a closed TP (Step 1). The three vertical edges of the TP are of double-stranded structure, which guarantees nanoassembly with a rigid 3D structure. The six horizontal edges of the top and bottom faces of the DNA nanostructure provide the hybridization sites for the combination of different functional sensor modules together. In this case, the three single-strand edges on the top face were used for the fixation of different capture sequences for target recognition. A diethylaminocoumarin (DEA)-labelled reference probe can be trapped on the bottom face of the TP, allowing for ratiometric self-calibration analysis (Step 2).31 The biotin sequence acts as an anchor module that holds these nanostructures onto the microbead surface to integrate them into devices for microbead assays (Step 3). Presence of the targets resulted in the formation of a sandwich hybridization structure, and strong fluorescent emission on the TP framework allows for multiplexed biomolecule detection (Step 4). Therefore, based on the well-defined 3D geometry and controlled assembly capability, the modular structure of the TP sensor provides a flexible, facile microbead sensing platform for ratiometric self-calibration imaging and multiplexed biomolecule detection. The detection of sequence-specific nucleic acids associated with human diseases has attracted much attention for early diagnosis and use in biological research.32 For instant, human immune deficiency virus (HIV), Ebola virus (EV) and human papillomavirus (HPV) are difficult to treat and highly infective, while coinfection with these viruses is considered to be deadly.33 In our system, oligonucleotide sequences of 30–33 bases associated with HIV, EV and HPV were chosen as representative targets.

Materials and Reagents. Streptavidin-modified sepharose beads (STV-MBs, 34 µm; biotin binding capacity, >300 nmol/mL medium) were purchased from GE Healthcare BioSciences (Sweden). Synthetic oligonucleotides were ordered from Shanghai Sangon Biological Engineering Technology (China), and TaKaRa Biotechnology Co., Ltd (Dalian, China). The details of the sequences used in this work are listed in Table S1. Other chemicals, if not specified here, were all commercially available and used without further purification. All aqueous solutions were prepared exclusively with Milli-Q water (18.2 MΩ). Construction of DNA TP nanostructure. First, equimolar amounts of the three scaffold clip sequences (Clip 1–3, 660 nM) were combined in TAMg buffer (45 mM Tris Acetate, 12.5 mM Mg(CH3COO)2, pH 8.0) followed by the annealing protocol using an automated PCR thermocycler: 95 °C for 5 min, 65 °C for 30 min, 50 °C for 30 min, 37 °C for 30 min, 22 °C for 30 min, and 4 °C for 55 min. The final concentration of the TP was estimated to be 220 nM. The TP was stored at 4 °C in the dark as a stock solution for further use. The three capture sequences, reference probe, and anchor sequence with 6 µM were loaded, in order, onto the TP (1 equiv/TP) using an automated PCR thermocycler: 50 °C for 30 min, 37 °C for 30 min, 22 °C for 30 min, and 4 °C for 55 min. It was supposed that the product yield was almost 100%, the final assembly of these functional strands combined with TP was designated TP5 (~186 nM) and was stored at 4 °C (Fig. S1). Microbead immobilization and imaging. The agarose microbeads were first washed with buffer solution to remove the ethanol solution. For microbead immobilization, 186 nM TP5 (960 µL) was incubated with 20 µL of microbead suspension (4.9 ×106 counts/mL) in TAMg buffer for 2 h with shaking at room temperature, resulting in the formation of TP5immobilized microbeads (TP5-MBs). The TP5-MBs were washed with TAMg buffer solution and stored at 4 °C for further use.

EXPERIMENTAL SECTION


Page 3 of 8

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 EV report probe was used as a model for single-target microbead imaging and ratiometric analysis. EV target with different concentrations was added to 50 nM TP5-MBs suspension containing the EV report probe (50 nM). After incubation in a rocker shaker at 25 rpm for 2 h at room temperature, the suspension was separated by centrifugation and redispersed in TAMg buffer solution. The fluorescence of the microbeads was recorded using a florescence microscope. About more than 30 microbeads were chose for simultaneous intensitybased analysis and ratiometric analysis at each sample. The linear equation was fitted and the limit of detection (LOD) was calculated from the signal that was equivalent to 3 times the standard deviation of the blanks (target 0 nM). For simultaneous detec-

tion of multiplex DNA targets, the HIV, EV and HPV targets were added to the 50 nM TP5-MBs suspension containing 50 nM FAM, ROX and Cy5 report probes and incubated for 2 h at 25 rpm in TAMg buffer at room temperature. After washing, the DEA, FAM, ROX and Cy5 fluorescence from the microbeads was recorded at different fluorescent channels using a fluorescence microscope. Instrument and Characterization. Sample dissolved in TAMg buffer were quantified using a Shimadzu UV-2600 spectrophotometer. Fluorescent spectra were measured using a Hitachi F4500 fluorescence spectrophotometer. Polyacrylamide gel electrophoresis (PAGE) was carried out in TAMg buffer at 120 V (constant voltage) for 2–3 h using a Bio-Rad electrophoresis apparatus (Bio-Rad Laboratories, Inc., Hercules, CA, USA). Gels were visualized using a Tanon-2500R gel imaging system (Shanghai, China). For dynamic light scattering (DLS), all measurements were performed using a Malvern Zetasizer Nano-ZS instrument equipped with a 633-nm laser. Samples were injected into a disposable zeta cuvette, and experiments were carried out at 25°C with a scattering angle of 175°C. Fluorescent imaging microscopy. Optical and fluorescence measurements were performed using a Nikon TE-2000U inverted microscope with a 20× objective. Fluorescence images were obtained using the following optical settings. The DAPI5060C filter cube (Semrock) was used for DEA. The FITC3540B filter cube (Semrock) was used for FAM. The TxRed4040C filter cube (Semrock) was used for ROX, and the Cy54040A filter cube (Semrock) was used for Cy5. All optical settings were dependent on the fluorescent tag(s) selection (Fig. S2). The fluorescence images were captured on a C910013 EMCCD (Hamamatsu). Ratiometric imaging analysis was conducted in Image-Pro Plus. The signal fluorescence images and reference fluorescence images were first corrected for background. The fluorescence intensities of the microbeads in the image then were calculated using Image-Pro Plus. Ratiometric data were calculated as the ratio between the signal and the corresponding reference fluorescence intensity. Atomic force microscopy (AFM). The diluted sample was pipetted onto freshly cleaved mica. After 20 min, the mica surface was washed several times with purified water and gently blown dry with nitrogen gas. The prepared sample was scanned using ScanAsyst-air tips in ScanAsyst Imaging Mode on a Multimode 8 Atomic Force Microscope with a NanoScope V controller (USA). Transmission electron microscopy (TEM). TEM was conducted using a Tecnai G2 F20 at 180 kV. Typically, droplets of the sample were applied to a carbon-film covered 200-mesh

grid (G2020, Agar Scientific, UK). The supernatant fluid was removed with filter paper after 1 min. Uranyl acetate (2%) then was dropped onto the sample for staining. Finally, the samples were dried in air under ambient conditions.

RESULTS AND DISCUSSION To form the DNA TP nanostructure in the formation of three-clip products C1-C2-C3, three 96-base clips sequences were combined and annealed at a temperature range of 95°C– 4°C over 3 h. Assembled structures were characterized by native PAGE (Fig. 2A). Successive additions of the three component strands generated bands of reduced mobility, indicating that successful hybridization of each component (lane 1–3) produced the 3D TP nanostructure in excellent yield (lane 3). TP (288 nt) mobility corresponded to the marker position between 140 and 160 bp. Some byproducts at higher molecular weight were probably due to the formation of polygon nanoprisms. In fact, the mixture of three clips C1, C2, C3 may result in the formation of six-clip byproducts C1-C2-C3C1-C2-C3 in the form of hexagonal prism, twelve-clip byproducts C1-C2-C3-C1-C2-C3-C1-C2-C3 in the form of nonagonprism, and so on. Those forms would have higher molecular weight, and migrated more slowly than TP. With heating to 95°C followed by slow cooling over 3 h, the three component strands annealed to yield quantitative formation of TP (lane 4). The melting curve and its negative first derivative showed a broad thermal denaturation range, with melting temperatures (Tm) of 43°C (Fig. 2B), consistent with the previous results.34 The size of the rigid TP construction was estimated to be 10.4 nm if the length of the 20-base TP edge was approximately 6.8 nm (Fig. S3). Analysis of the TP by dynamic light scattering (DLS) gave a hydrodynamic diameter of 13.8 ± 4.2 nm with a near monodisperse particle population (Fig. S4), which was slightly larger than the theoretical value. However, atomic force microscopy (AFM) indicated the TP height as 1.2 ± 0.5 nm (Fig. 2C, D), smaller than the theoretical value of 10.4 nm. This difference may be caused by the sample preparation, in which deposition of TP on mica and the air-dried process can result in TP collapse and distortion, that was previously reported.32 This size was also validated by the uranyl acetate-stained TEM images, indicating a TP diameter of 2.9 ± 0.5 nm (Fig S5).

3



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 2. Characterization of the TP nanostructure. (A) Native PAGE analysis of TP assembly. Native PAGE (6%; all samples assembled in TAMg). Lanes 1 to 3 are RT additions of component strands, and lane 4 is the thermocycled final product. Lane 1, C1; Lane 2, C1 + C2; Lane 3, C1 + C2 + C3; Lane 4, C1 + C2 + C3 (thermocycled); Lane 5, marker. (B) Negative first derivative of the melting curve of TP. Inset: the melting curve of TP. The fluorescence intensity of the TP decreased with increasing temperature. (C) AFM imaging of the TP nanostructure. Scale bar: 100 nm. (D) Line profile of a cross section in AFM imaging (C).

This TP cage possesses 6 single-strand binding regions with different 20-base sequences that were exploited to assemble different sensing modules together into a single TP to construct the sensor system (Fig S1). Fig. 3A demonstrates the stepwise assembly of three capture sequences and a reference probe on the TP, with corresponding decreases in gel mobility from lane 1 to lane 5. The equal decrease in migration from lane 1 to lane 4 demonstrates the immobilization of three similar capture sequences for HIV, EV and HPV. The assemblies in lane 4 are designated TP3, resulting from the combining of TP with three capture sequences. Binding of the reference probe to TP3 yielded the TP4 nanostructures in lane 5. Furthermore, loading of the anchor sequence onto TP4 yielded TP5, as shown in Fig. 3B. After incubation of the TP5 probe with STV-MBs microbeads for 2 h, intense blue fluorescence was observed from the microbeads (Fig. 3D). In contrast, STV-MBs incubated with TP4 without the anchor sequence showed almost no fluorescence (Fig. 3C), suggesting that the anchor sequence was crucial for immobilization of the TP scaffold onto STV-MBs. The immobilization capacity was effectively adjusted by controlling the mixture ratio of TP5 and the microbeads; when saturated about 7.9×108 units of TP5 were anchored onto each STV-MB, on average (Fig S6). Therefore, in this case, the single-strand sequence on the bottom face of the TP acted as an anchor binding site in the immobilization module of the TP5-MBs system. The immobilization module provided a broad platform for the utilization of the DNA nanostructures in the solid phase assay, such as SPR, microfluidic chip and microbead assay.

Figure 3. (A) Native PAGE analysis of the assembly of different sensing modules on TPs. Lanes 1 to 4 indicate the stepwise assembly of TPs. Lane 1, TP; Lane 2, TP1= TP + capture HIV; Lane 3, TP2= TP1 + capture EV; Lane 4, TP3= TP2 + capture HBV; Lane 5, TP4= TP3 + reference sequence. (B) Native PAGE analysis of the assembly of anchor sequences onto TPs. Lane 1, TP4; Lane 2, TP5= TP4 + anchor sequence. (C, D) Bright and fluorescent images of STV-MBs after incubation with TP5 and TP4. Scale bars: 10 µm.

Page 4 of 8

The immobilization of TP nanostructures onto microbeads could be reversibly adjusted through strand displacement. In our design, the anchor sequence designed to hybridize to the single-strand regions of the TP consisted of a complementary 20-base region followed by a T10 extension and chemical modification at the 5’ end (Fig. S7). The T10 extension served as a toehold initiation point for strand displacement of the 30mer DNA conjugates from the TP5 scaffold.35 In this way, addition of the complementary sequence (anchor displacement sequence) selectively displaced the anchor sequence from the nanoprism, thereby displacing the TPs from the microbeads. Therefore, the binding and removal of the anchor sequence would result in the reversible immobilization of the TP nanostructures on the microbeads. The TP5 probe could be exploited for the single microbead fluorescent assay. Although microbead assays are widely used in functional genomics experiments and protein screening, uneven intensity distribution and different probe concentrations on microbeads can cause imaging deviations.36 Compared to intensity-based sensing, ratiometric sensing using the built-in correction of two different emission bands is more ideal for practical applications because differences in microbead and probe concentrations can be corrected.37 Here, the immobilization of a reference probe onto the TP framework allows for ratiometric assay. In our microbead assay, the DEA-labelled reference probe and ROX-labelled EV report probe were combined on the TP nanostructure as a representative ratiometric sensing platform for microbead assay. A 10 nM EV target was added to the 50 nM TP5-MBs reaction system containing a 50 nM EV report probe. After incubation for 2 h, the microbeads were washed for fluorescence imaging. Both ROX and DEA fluorescence on the microbeads possessed strong yellow and blue fluorescence, and the fluorescence was well distinguished using different fluorescent channels (Fig. 4A, B). Even for the single microbead, the fluorescence intensity showed large variance at the centre and periphery of the microbeads caused by differing microbead refractions. Quantitative analysis of the plot profile for the crossing line, illustrated as ‘M’ letter morphology, and the fluorescence at the periphery was almost twice the fluorescent intensity as that in the centre (Fig 4D, E, microbead ①). After ratiometric data processing, a representative ratiometric image (Fig 4C, F) demonstrated the variance at the centre and periphery of the microbeads was largely reduced and the offset of the uneven fluorescent intensity indicated a single microbead with uniform intensity distribution. Although the color variance at the centre and periphery still existed in a certain degree (Fig. 4C), the variance of intensity at the center and periphery of the microbeads was greatly decreased. The reliability of the microbead imaging was further investigated. It was demonstrated that after ratiometric analysis, the error from the uneven intensity distribution of the single microbead was greatly reduced, and the relative standard deviation (RSD) decreased from 0.57 to 0.20 (Fig. S8). Therefore, TP nanostructures based microbeads immobilization provides a more reliable microbead analysis approach. The different TP5 probe concentrations immobilized on microbeads also contribute to the imaging deviation observed in the intensity-based analysis. This can be overcame in the ratiometric analysis, since each TP5 framework had the possibility to bind one of the target molecules and a reference probe,

4


Page 5 of 8

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


and thus the ratiometric signal was independent of the TP5 nanostructure concentration immobilized on the microbeads. As demonstrated in Fig. 4(H-N), the 50 nM EV target was added to two TP5-MBs mixtures with differing TP5 concentrations on the microbeads. Microbead ② and ③ differed greatly in intensity in both the blue and yellow channel fluorescence (Fig 4H, I). The fluorescence of microbead ② was almost 1.5 times the intensity of microbead ③. We attributed this difference to the differing concentrations of TP5 immobilized on the microbeads. After the ratiometric data processing, the offset of the uneven fluorescent intensity resulted in two microbeads with uniform intensity distribution (Fig 4J, N). In this case, the error from concentration differences in TP5 immobilization was greatly reduced. Therefore, ratiometric analysis using a reference probe was more reliable than that with intensity-based probes for the detection of DNA targets. The modular TP probe provides a reliable single-microbead imaging platform for quantitative analysis.

The performance of quantitative ratiometric analysis for EV target detection was further investigated by fluorescence imaging of single microbeads upon the addition of varying concentrations of EV target in 50 nM TP5-MBs solution containing a 50 nM EVreport probe. A large number of TP5 probes were modified on the microbeads through a biotin anchor sequence, and the blue fluorescence of the microbeads differed slightly between samples (Fig 5). With a gradual increase in EV concentration, the fluorescence intensity of the ROX unit (yellow channel) on the microbeads gradually increased. The ratiometric images were calculated from the ratio of ROX to DEA fluorescence. At the range of 0–50 nM, the ratiometric results showed a similar fluorescent response trend to that of ROX fluorescence, indicating that ratiometric analysis was effective for quantitative detection. More important, with regard to ratiometric analysis, single microbead analysis demonstrated an even intensity distribution with a small relative standard deviation, which resulted in greater accuracy in quantitative analysis. A limit of detection (LOD) down to 0.75 nM was achieved for the EV target. This result indicates that this sensing system can be used effectively to measure targets with a linear calibration curve in the concentration range from 0 to 40 nM (Fig. S9).

Figure 5. Quantitative intensity-based and ratiometric analysis of a DNA target on a single microbead. Fluorescence imaging of the microbead upon addition of varying concentrations of EV targets in 50 nM TP5-MB solution containing a 50 nM ROX probe. DEA, blue fluorescence from reference probe; ROX, yellow fluorescence from ROX labelled EV report probe; Ratio, ratiometric analysis of ROX fluorescence to DEA fluorescence. Figure 4. (A–F) Ratiometric fluorescent imaging of TP5-MBs for sensing the EV target. Fluorescence image of (A) DEA fluorescence and (B) ROX fluorescence from microbeads containing a 50 nM ROX report probe upon addition of 10 nM ROX target. (C) Ratiometric images of ROX fluorescence to DEA fluorescence. Scale bar: 50 µm. (D, E, F) Line profile of the cross section line for microbead ① from A, B and C, respectively. (H-N) Ratiometric fluorescent imaging of mixed TP5-MBs for sensing the EV target. Two types of TP5-MBs modified with 7.9×108 units of TP probe and 5.0 ×108 units of TP probe per microbead were mixed together. Fluorescence image of (H) DEA fluorescence and (I) ROX fluorescence from the microbeads containing a 50 nM ROX report probe upon addition of 50 nM ROX targets. (J) Ratiometric images of ROX fluorescence to DEA fluorescence. Scale bar: 50 µm. (L, M N) Line profile of the cross section line for microbead ② and ③ from H, I and J, respectively.

With regard to the TP5 nanostructure, the three single-strand regions on the top face provide three recognition sites, conferring TP5-MBs with the potential for positioning multiplex capture sequences for simultaneous detection. Different target samples for EV, HIV and HPV were added to the 50 nM TP5MBs sensing system containing 50 nM HIV, EV and HPV report probes (Fig. S10). This method involves the formation of three sandwich hybridization structures labelled with different fluorescent molecules. However, when different fluorophores are attached together, fluorescence resonance energy transfer (FRET) between different fluorophores may occur. Here, the efficiency of FRET from FAM to ROX, and from ROX to Cy5 were estimated to be 1.8×10-9, and 5.1×10-9, respectively (Supporting Information). Therefore, no obvious FRET occurred in this sandwich structure, which was validated by the observation that no obvious fluorescence fluctuation

5



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

occurred after the fluorescent report probes were attached to the TP nanostructure (Fig S11). This result also demonstrates that this 3D geometrically well-defined TP provides a versatile framework for spatial orientation, allowing positioning of multiplex sensing modules that will not interfere with each other. In this system, the fluorescence from different channels can be well discriminated. As shown in Fig. 6, when the three targets were present at the same time (Sample 1), the fluorescence signals of all three channels were ‘ON’. On the other hand, two out of three channels showed detectable signals when two targets coexisted (Samples 2–4). When only one of the targets was present (Sample 5–7), the fluorescence signal of the corresponding channel was ‘ON’. We also challenged the specificity of this microbead imaging system using no targets (Sample 8). No detectable signal from the three channels was observed. Different targets resulted in differential patternsensing spectra, through which we obtained unique ratiometric finger-print pattern arrays with unique fluorescence change features that could be used to discriminate between different targets. Therefore, this TP nanostructure imaging system provides a flexible and versatile microbead ratiometric sensing platform that shows great promise for the discrimination of multiplex targets.

Figure 6. TP5-MBs for fingerprint-like multiplex pattern sensing. Fluorescence imaging of the microbead upon addition of 30 nM of EV, HIV or HPV targets in 50 nM TP5-MBs solution containing 50 nM ROX probe, FAM probe, or Cy5 probe. (A) Fluorescence imaging of microbeads at different fluorescent channels. Blue fluorescence from DEA-labelled reference probe; Yellow fluorescence from ROX-labelled HPV report probe; Green fluorescence from FAM-labelled HIV report probe; Red fluorescence from Cy5-labelled HPV report probe. (B) Fingerprint-like patternsensing for multiplex target discrimination based on ratiometric analysis.

It is worth noting that the TP nanostructures could be regenerated and the sandwich hybrid structure for the multiplex detection could be disassembled through sequence displacement (Fig 7). Addition of the partial complementary sequence (HPV displacement sequence) to the HPV capture sequence selectively displaced the HPV target sequence from the TP nanostructures. Subsequently, the red fluorescence from the Cy5 HPV report probe disappeared. Using the same strategy, the sandwich hybrid structures for the HIV and EV detection were also disassembled, and there was no yellow or green fluorescence from the HIV or EV report probes in microbead

Page 6 of 8

imaging. Disassembly of the sandwich hybrid structure provides the option to regenerate the binding sites on the TP top face. Therefore, this displacement assembly approach allows for the regeneration of the TP sensor and also further incorporation of diverse functionalized ligands onto the TP nanostructure, expanding the sensing applications.

Figure 7. Sequence displacement strategy for TP nanostructure regeneration. (A) Addition of the HPV displacement sequence selectively displaces the HPV sandwich hybrid structures from the TP (Step 1, lanes 1–2). Subsequent addition of the HIV displacement sequence selectively displaces the HIV sandwich hybrid structures from the TP (Step 2, lanes 2–3). Subsequent addition of the EV displacement sequence selectively displaces the EV sandwich hybrid structures from the TP (Step 3, lanes 3–4). Accordingly, the red fluorescence, green fluorescence, and yellow fluorescence signals from the Cy5-labelled HPV report probe, FAM labelled HIV report probe and FOX labelled EV report probe, respectively, disappeared. Based on this sequence displacement strategy, the combination sites in the TP nanostructure were regenerated. (B) Detailed sequence displacement process for Step 2.

CONCLUSION In summary, we have integrated different functional modules into a 3D DNA TP nanoassemblies and used the multiple single-strand region to reconfigure this nanostructure, enabling ratiometric and multiplexed biomolecule detection on a single microbead. Taking the TP framework as a flexible and modular platform, the anchor module, the reference probe and multiple capture sequences were integrated together on this single TP system. This approach significantly simplifies the design and regeneration of diverse analytical devices by allowing for the alteration of functional module strands while retaining the DNA nanostructure. Ratiometric analysis indicates that the imaging deviation caused by differential distribution on single microbead and the different TP5 concentrations immobilized on microbeads was greatly reduced. Through the application of a finger print-like pattern array, we are able to use these fluorescence responses to discriminate between different tar-

6


Page 7 of 8

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


gets in a simple and efficient fashion. Therefore, the modular 3D DNA nanostructure shows great promise for biomedical imaging and high-throughput microbead analysis.

ASSOCIATED CONTENT Supporting Information DNA sequences, absorption and fluorescent spectra, TEM imaging, gel electrophoresis, DLS, and LOD calculation. The Supporting Information is available free of charge on the ACS Publications website.

AUTHOR INFORMATION Corresponding Author *Tel.: 86-731-88821566. Fax: 86-731-88821566. E-mail: [email protected]; [email protected].

Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT This work was supported by the National Natural Science Foundation of China (21190040, 21575037, 21675046).

REFERENCES (1) Woo, S.; Rothemund, P. W. Nat. Commun. 2014, 5, 4889. (2) Lu, N.; Pei, H.; Ge, Z.; Simmons, C. R.; Yan, H.; Fan, C. J. Am. Chem. Soc. 2012, 134, 13148-13151. (3) Li, W.; Yang, Y.; Yan, H.; Liu, Y. Nano Lett 2013, 13, 2980-2988. (4) Genot, A. J.; Bath, J.; Turberfield, A. J. J. Am. Chem. Soc. 2011, 133, 20080-20083. (5) Chen, Y. J.; Groves, B.; Muscat, R. A.; Seelig, G. Nat. Nanotechnol. 2015, 10, 748-760. (6) Chen, G.; Liu, D.; He, C.; Gannett, T. R.; Lin, W.; Weizmann, Y. J. Am. Chem. Soc. 2015, 137, 3844-3851. (7) Wan, L.; Chen, Q.; Liu, J.; Yang, X.; Huang, J.; Li, L.; Guo, X.; Zhang, J.; Wang, K. Biomacromolecules 2016, 17, 15431550. (8) Xu, L.; Yan, W.; Ma, W.; Kuang, H.; Wu, X.; Liu, L.; Zhao, Y.; Wang, L.; Xu, C. Adv. Mater. 2015, 27, 1706-1711. (9) Jiang, Q.; Shi, Y.; Zhang, Q.; Li, N.; Zhan, P.; Song, L.; Dai, L.; Tian, J.; Du, Y.; Cheng, Z.; Ding, B. Small 2015, 11, 51345141. (10) Fakhoury, J. J.; McLaughlin, C. K.; Edwardson, T. W.; Conway, J. W.; Sleiman, H. F. Biomacromolecules 2014, 15, 276282. (11) Tay, C. Y.; Yuan, L.; Leong, D. T. ACS Nano 2015, 9, 5609-5617. (12) Kong, R. M.; Zhang, X. B.; Chen, Z.; Tan, W. Small 2011, 7, 2428-2436. (13) Lin, M.; Wang, J.; Zhou, G.; Wang, J.; Wu, N.; Lu, J.; Gao, J.; Chen, X.; Shi, J.; Zuo, X.; Fan, C. Angew. Chem. 2015, 54, 21512155. (14) Liu, Y.; Lin, C.; Li, H.; Yan, H. Angew. Chem. Int. Ed. 2005, 44, 4333-4338. (15) Liu, Z.; Tian, C.; Yu, J.; Li, Y.; Jiang, W.; Mao, C. J. Am. Chem. Soc. 2015, 137, 1730-1733. (16) Seeman, N. C. Mol. Biotechnol. 2007, 37, 246-257. (17) Mei, Q.; Wei, X.; Su, F.; Liu, Y.; Youngbull, C.; Johnson, R.; Lindsay, S.; Yan, H.; Meldrum, D. Nano Lett. 2011, 11, 14771482. (18) Sannohe, Y.; Endo, M.; Katsuda, Y.; Hidaka, K.; Sugiyama, H. J. Am. Chem. Soc. 2010, 132, 16311-16313.

(19) Selmi, D. N.; Adamson, R. J.; Attrill, H.; Goddard, A. D.; Gil-bert, R. J.; Watts, A.; Turberfield, A. J. Nano Lett. 2011, 11, 657660. (20) Cohen, J. D.; And, J. P. S.; Dervan, P. B. J. Am. Chem. Soc. 2008, 130, 402-403. (21) Goodman, R. P.; Heilemann, M.; Doose, S.; Erben, C. M.; Ka-panidis, A. N.; Turberfield, A. J. Nat. Nanotechnol. 2008, 3, 9396. (22) Wen, Y.; Pei, H.; Shen, Y.; Xi, J.; Lin, M.; Lu, N.; Shen, X.; Li, J.; Fan, C. Sci. Rep. 2012, 2. (23) Wen, Y.; Pei, H.; Wan, Y.; Su, Y.; Huang, Q.; Song, S.; Fan, C. Anal. Chem. 2011, 83, 7418-7423. (24) Chen, X.; Hong, C.-Y.; Lin, Y.-H.; Chen, J.-H.; Chen, G.N.; Yang, H.-H. Anal. Chem. 2012, 84, 8277-8283. (25) Conway, J. W.; Madwar, C.; Edwardson, T. G.; McLaughlin, C. K.; Fahkoury, J.; Lennox, R. B.; Sleiman, H. F. J. Am. Chem. Soc. 2014, 136, 12987-12997. (26) Edwardson, T.; Lau, K.; Bousmail, D.; Serpell, C.; Sleiman, H. F. Nat. Chem. 2016, 8, 162–170. (27) He, K.; Li, Y.; Xiang, B.; Zhao, P.; Hu, Y.; Huang, Y.; Li, W.; Nie, Z.; Yao, S. Chem. Sci. 2015, 6, 3556-3564. (28) Dai, Z.; Gao, Q.; Cheung, M.C.; Leung, H.M.; Lau, T.C.K.; Sleiman, H. F.; Lai,K.W.C.; Lo, P.K.. Nanoscale, 2016,8, 1829118295. (29) Stoeva, S. I.; Lee, J. S.; Thaxton, C. S.; Mirkin, C. A. Angew. Chem. 2006, 45, 3303-3306. (30) Song, S.; Liang, Z.; Zhang, J.; Wang, L.; Li, G.; Fan, C. Angew. Chem. 2009, 48, 8670-8674. (31) He, L.; Yang, X.; Zhao, F.; Wang, K.; Wang, Q.; Liu, J.; Huang, J.; Li, W.; Yang, M. Anal. Chem. 2015, 87, 2459-2465. (32) Sun, S.; Yao, H.; Zhang, F.; Zhu, J. Chem. Sci. 2015, 6, 930-934. (33) Zhang, S.; Han, G.; Xing, Z.; Zhang, S.; Zhang, X. Anal. Chem. 2014, 86, 3541-3547. (34) Conway, J. W.; McLaughlin, C. K.; Castor, K. J.; Sleiman, H. Chem. Comm. 2013, 49, 1172-1174. (35) Zhang, D. Y.; Seelig, G. Nat. Chem. 2011, 3, 103-113. (36) Theilacker, N.; Roller, E.E.; Barbee, K. D.; Franzreb, M.; Huang, X. J. R. Soc. Interface 2011, 8, 1104–1113. (37) Huang, J.; Ying, L.; Yang, X.; Yang, Y.; Quan, K.; Wang, H.; Xie, N.; Ou, M.; Zhou, Q.; Wang, K. Anal. Chem. 2015, 87, 87248731.

7



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 8 of 8

Table of Contents (TOC)

8


Design of a Modular DNA Triangular-Prism Sensor Enabling

Recommend Documents