Fluorescence Resonance Energy Transfer-Based DNA Nanoprism


Sep 21, 2017 - Here, combining the excellent biological properties of DNA nanostructures with low background split aptamers, we developed a functional...
3 downloads 19 Views 935KB Size


Subscriber access provided by University of Sussex Library

Article

Fluorescence Resonance Energy Transfer-Based DNA nanoprism with a Split Aptamer for ATP sensing in living cells Xiaofang Zheng, Ruizi Peng, Xi Jiang, Yaya Wang, Shuai Xu, Guoliang Ke, Ting Fu, Qiaoling Liu, Shuang-Yan Huan, and Xiaobing Zhang Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b02763 • Publication Date (Web): 21 Sep 2017 Downloaded from http://pubs.acs.org on September 22, 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 22

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

Fluorescence Resonance Energy Transfer-Based DNA nanoprism with a Split Aptamer for ATP sensing in living cells

Xiaofang Zheng, Ruizi Peng, Xi Jiang, Yaya Wang, Shuai Xu, Guoliang Ke, Ting Fu, Qiaoling Liu, Shuangyan Huan*, Xiaobing Zhang

Molecular Sciences and Biomedicine Laboratory, State Key Laboratory of Chemo/Biosensing and Chemometrics, College of Chemistry and Chemical Engineering, College of Biology, Hunan University, Changsha 410082, P. R. China.

*Email: [email protected]; Tel: 86-731- 88821632

ACS Paragon Plus Environment

Analytical Chemistry

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

ABSTRACT: We have developed a DNA nanoprobe for adenosine triphosphate (ATP) sensing in living cells, based on the split aptamer and the DNA triangular prism (TP). In which nucleic acid aptamer was split into two fragments, the stem of the split aptamer was respectively labeled donor and acceptor fluorophores that underwent an fluorescence resonance energy transfer if two ATP molecules were bound as target molecule to the recognition module. Hence, ATP as a target induced the self-assembly of split aptamer fragments and thereby brought the dual fluorophores into close proximity for high FRET efficiency. In the vitro assay, an almost 5-fold increase in FA/FD signal was observed, the fluorescence emission ratio was found to be linear with the concentration of ATP in the range of 0.03-2 mM, and the nanoprobe was highly selective toward ATP. For the strong protecting capability to nucleic acids from enzymatic cleavage and the excellent biocompatibility of the TP, the DNA TP nanoprobe exhibited high cellular permeability, fast response, and successfully realized “FRET off” to “FRET on” sensing of ATP in living cells. Moreover, the intracellular imaging experiments indicated that the DNA TP nanoprobe could effectively detection ATP and distinguish among changes of ATP levels in living cells. More importantly, using of the split aptamer and the “FRET off” to “FRET on” sensing mechanism could efficiently avoid false-positive signals. This design provided a strategy to develop biosensors based on the DNA nanostructures for intracellular molecules analysis.

ACS Paragon Plus Environment

Page 2 of 22

Page 3 of 22

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

INTRODUCTION In living organisms, adenosine triphosphate (ATP) as the major energy source powers, regulates many biological pathways and extracellular signaling mediator in many biological processes.1-3 ATP level is an indicator for many diseases such as hypoxia, ischemia, Parkinson’s disease, and some malignant tumors.4-6 Various strategies have been used for ATP detection, such as electrophoresis (EP), isotope tracing method (ITM), chemiluminescence (CL), and so on. However, these methods are time-consuming, require tedious sample preparations, and even cannot be used in ATP imaging in living cells, which has prompted the search of alternative approaches for ATP detection in living cells. With quick development of biotechnologies, aptamers have attracted widespread attention for their high affinity, high specificity, and real-time and in situ monitoring capability.7,8 Aptamers have been widely used in analyzing small molecules, biomacromolecules, and organisms.9-11 But there are several problems need to be addressed when using nucleic acid-based fluorescent probes for the detection and imaging of ATP in living cells. The first problem is steric hindrance of excessive bases, because aptamers with the target is in essence the combination of bases and target sites. The second is effective delivery of probes into the cell. Single nucleic acid probe exists poor cellular internalization and easily enzymes degradation. Lastly, the “always on” sensing mechanism, which lack target-activatable nature will inevitably result in high-background and low-contrast image, delayed diagnosis and toxicity to normal tissues.12-14 To circumvent these problems, Stojanovic et al. notably developed a split aptamer-based strategy, in which nucleic acid aptamers were split into two fragments and they can specifically form a ternary complex in the presence of target. Owing to the lack of secondary structures between two separate oligonucleotides, almost no false-positive or nonspecific signals was yielded.15 This strategy has been widely adopted for detection of various targets, especial small molecules, using different

transduction

methods

including

colorimetric,

fluorescence

and

electrochemical techniques,16-23 But currently, utilizing split aptamer is still in the

ACS Paragon Plus Environment

Analytical Chemistry

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

infancy stage and studies are focused on simple improvement of sensitivity. Stability is still a problem and false-positive signals cannot be entirely avoided. DNA nanostructure provide a biocompatible tool for delivering these nucleic acid probe. Among various biosensors, DNA-based nanostructures exhibit unique merits. Firstly, DNA molecules are inherently biocompatible and biodegradable.24,25 Secondly, DNA aptamers can fold into well-defined shapes to recognize targets with high selectivity and affinity, which ranging from small inorganic and organic molecules to macromolecules, or even cells.26 Moreover, DNA molecules are not merely the genetic biopolymer carrier but also widespread building block in materials science.27-31 Recently, various static DNA nanostructures with unmatched precision like 2D and 3D DNA structures based-on tile motifs, discrete DNA origami nanostructures and DNA brick architectures have been constructed via molecular self-assembly.31-33 In particular, DNA brick self-assemblies have attracted extensive concern because it utilized a DNA-minimal approach to create highly functional objects including DNA nanotubes and DNA cages, such as tetrahedron, triangular prism, cube, pentagonal prism, etc. DNA cages can encapsulate hydrophobic micelles for therapeutic drugs delivery,34-36 be modified with gold nanoparticles,37 or even function as useful tools to probe the size and shape dependence of nucleic acid delivery.38,39 Numerous of DNA nanostructures have been reported that can cause a specific biological response in vitro or in cells. For example, Sleiman et al. assembled a trigger-responsive siRNA-encapsulating DNA cage and investigated its target response in a complex extracellular environment.30 Liu et al. have realized multiplexed biomolecule detection on a single microbead by designing a DNA triangular-prism sensor.40 He et al. developed a DNA tetrahedron nanotweezer nanoprobe for imaging tumor-related mRNA in living cells based on the FRET.41 Here, combining the excellent biological properties of DNA nanostructures with low background split aptamers, we developed a functional nanomechanical DNA triangular prism device that encapsulating a split aptamer.42-45 The stem of the split aptamer was respectively labeled donor and acceptor fluorophores that underwent a

ACS Paragon Plus Environment

Page 4 of 22

Page 5 of 22

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

fluorescence resonance energy transfer (FRET) if two ATP molecules were bound as target molecule to the recognition module. Scheme 1 illustrates the principle of DNA TP Nanoprobe for ATP sensing in Living Cells. In absence of the target the two gating strands were spatially separated, the respectively labeled donor and acceptor fluorophores were separated, and FRET was in the “off” state. In the presence of the marker, the two gating strands were from open to closed state, bringing the dual fluorophores into close proximity and leading to high FRET efficiency. Therefore, based on the DNA nanostructures controlled assembly capability, the modular structure of the TP nanosensor provides a flexible, facile microbead sensing platform for detecting ATP in living cells and in vivo with high sensitivity and image resolution, which leads to versatile functions in cellular sensing and imaging of the predicting biomarkers such as mRNAs and microRNAs.

EXPERIMENTAL SECTION Materials and Reagents. Sequences of Oligonucleotides used in this work are listed in Table S1, the materials for DNA synthesis were purchased from Glen Research (Sterling, VA). Adenosine triphosphate (ATP), cytidine triphosphate (CTP), uridine triphosphate (UTP), guanosine triphosphate (GTP) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Stains-All was purchased from Sigma-Aldrich Co., Ltd. The 20 bp DNA ladder was purchased from TaKaRa Biotechnology Co., Ltd. (Dalian, China). 1×TAE/Mg2+ buffer (20 mM Tris-acetic acid, 2 mM EDTA and 12.5 mM magnesium acetate, balanced to pH 7.5) was used for all self-assembled reactions. Complete medium (RPMI 1640) with 10% fetal bovine serum and penicillin (100 U/mL)-streptomycin (100 µg/mL) were used for cell culture. Washing buffer contained glucose (4.5 g/L) and MgCl2 (5 mM) in Dulbecco’s phosphate buffered saline (D-PBS; Gibco). Glass bottom cell culture dishes (Φ15 mm) were purchased from Nest Biotechnology Co., Ltd. (Wuxi, China). Other reagents from commercial suppliers were analytical grade and used without further purification. Ultrapure water

ACS Paragon Plus Environment

Analytical Chemistry

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

(electric resistance 18.2 MΩ) obtained through a Millipore Milli-Q water purification system (Billerica, MA, U.S.A.) was used throughout the experiments. Apparatus. Bio-Rad molecular imager (ChemiDoc XRS+ with lab imaging software), atomic force microscope (Veeco, Nanoscope IIIa), TC 10 automated cell counter (Bio-Rad), microplate reader (BioTek) and confocal laser-scanning microscope (Olympus, FV1000) were used in the experiments. Preparation of the DNA TP Nanoprobe. DNA TP was synthesized by mixing equimolar sequences (DNA strands were shown in Table S1) in 1×TAE/Mg2+ buffer (20 mM Tris-acetic acid, 2 mM EDTA and 12.5 mM magnesium acetate, balanced to pH 7.5). Followed by the annealing protocol using an automated polymerase chain reaction (PCR) thermal cycler: 95 °C for 5 min, 80 °C for 30 min, 79 °C for 30 min, 77 °C for 30 min and then gradually cooled to 4 °C in 1.5 h. The final concentration of the TP was estimated to be 1.5 µM, which was stored at 4 °C in the dark as a stock solution for further use. The two sequences of ATP-Apt1, ATP-Apt2 were modified on the TP (1 equiv/TP) by a slower cooling process from 85 to 25 °C over 4 hours. The final assembly of DNA TP Nanoprobe was stored at 4 °C (Figure S1). Electrophoresis Characterization. A 5% Native Polyacrylamide gel was prepared with 10.9 mL of Ultrapure water, 1.5 mL of 10×TAE/Mg2+, 2.5 mL of 30% Acryl-Bis, 0.11 mL of 10% APS and 0.010 mL of TEMED. DNA TP and DNA TP Nanoprobe were eventually quantified in a volume of 10 µL to give a desired concentration. Then, 2 µL of 6×loading buffer were added in each sample directly for electrophoresis experiments. Electrophoresis was carried out in 1×Tris-acetate-EDTA (TAE) buffer (40 mM Tris-HAc, and 1 mM EDTA, 12.5 mM Mg(Ac)2, pH 7.5) at 110 V for 1 h at room temperature. After stopping electrophoresis, gel was removed, and DNA bands were imaged and analyzed using a Bio-Rad molecular imager with imaging software under UV light. Moreover, to assess the stability of the DNA TP Nanoprobe, low (0.25 U/mL) and high (2.5 U/mL) concentration of Dnase I was respectively added to the sample solution containing 2 µM DNA TP Nanoprobe and

ACS Paragon Plus Environment

Page 6 of 22

Page 7 of 22

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

incubated with different time (0, 10, 20, 30, 40, 50, 60 min) at 37 °C. In order to further illustrate the stability of the DNA TP Nanoprobe, which was also mixed with FBS (10% v/v) in 1640 medium at 2 µM and incubated with different time (0, 1, 2, 3, 4, 6 h) at 37 °C. Then 12 µL of each sample including 2 µL of 6×loading buffer was added directly for electrophoresis experiments. 5% Native Polyacrylamide gel electrophoresis running at 110 V for about 30 min (4 °C) in 1× TAE buffer. DNA bands were imaged and analyzed using a Bio-Rad molecular imager with imaging software under UV light. AFM Imaging of DNA TP Nanoprobe. After 10 µL of DNA TP Nanoprobe were deposited 10 min on the surface of freshly cleaved mica, 5 µL of 30 mM Ni2+ were added to enhance the adhesion of samples. When the solution dried at room temperature, the mica was rinsed with Milli-Q water three times and dried in the nitrogen atmosphere. Atomic force microscopy of samples was observed on a Multimode 8 (Bruker/USA) using Scan Asystmode. In Vitro Detection of ATP. Different concentrations of ATP (0, 0.5, 1, 1.25, 1.5, 1.75, 2, 3, 4, and 5 mM) were added into 50 nM DNA TP Nanoprobe in the 1×TAE/Mg2+ buffer and the mixture was incubated at 37 °C for 1 h. The fluorescence emission spectra were recorded from 540 to 750 nm at an excitation wavelength of 525 nm in a 200 µL quartz cuvette. All experiments were repeated at least three times. Selectivity. Concentration of ATP, CTP, GTP, and UTP stock solution was added into containing 50 nM DNA TP Nanoprobe solution to reach a final concentration of 4 mM, respectively. After 30 min of incubation (37 °C), The fluorescence emission spectra were recorded from 540 to 750 nm at an excitation wavelength of 525 nm in a 200 µL quartz cuvette. All experiments were repeated at least three times. Cell Culture and Cytotoxicity. Hela cells (human cervical carcinoma) were obtained from ATCC (American Type Culture Collection, Manassas, VA, USA). Cell lines were cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum (FBS, heat inactivated) and penicillin(100 U/mL)-streptomycin (100 µg/mL) in

ACS Paragon Plus Environment

Analytical Chemistry

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

a cell culture incubator at 37 °C with 5% CO2. For adherent Hela cell lines, short-term (about 3 min) trypsin treatment was adopted to dissociate cells from culture flask or dish. Cell density was determined by using a hemocytometer. Cytotoxicity of the DNA TP nanoprobe was assessed with a standard MTS assay. Hela cells (1×104 cells/well) were cultured in 96-well microtiter plates and incubated at 37 °C in 5% CO2 for 24 h to ensure good adherence. Cells were incubated with different concentrations of DNA TP Nanoprobe (0, 50, 100, 200, and 400 nM) for 12 h at 37 °C in 5% CO2. The medium was directly removed. Each 20 µL of CellTiter reagent was added to 100 µL of fresh medium and cells were incubated at 37 °C in 5% CO2 for 20 min. A microplate reader was used to record the absorbance at 490 nm. Cell viability was determined according to the manufacturer’s description. Confocal Laser-scanning Microscopy Imaging. Hela Cells were seeded in a 15 mm confocal dish and incubated at 37 °C in 5% CO2 for 24 h. Then washing 3 times with DPBS, 500 µL of fresh cell growth medium supplemented with 50 nM of DNA TP nanoprobe was added in the dishes. After an incubation of 4 h, washing buffer was employed to wash the cells 3 times to remove the nanoprobes that were not uptaken by the cells. To monitor the ability of the nanoprobes for testing the levels of intracellular ATP. The Hela cells were treated with 10 µM oligomycin or with 5 mM Ca2+ for 30 min, then the nanoprobes-loaded cells were incubated at 37 °C for 4 h before imaging. The confocal microscope imaging of the cells was observed with a FV1000 confocal microscope (Olympus).

RESULTS AND DISCUSSION Design and Mechanism of The DNA TP Nanoprobe. Combining with the excellent chemical properties of DNA nanostructures, low background split aptamers and the incomparable advantages of “FRET-off” to “FRET-on” sensing mechanism in bioimaging, we designed a DNA TP nanoprobe which can enhance the stability and

ACS Paragon Plus Environment

Page 8 of 22

Page 9 of 22

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

efficiency of the aptamer. As shown in Scheme 1, DNA TP nanoprobe was prepared by a DNA-minimal nanoprism encapsulating a split aptamer which acts as two gates (Apt1 and Apt2 were modified with Cy3 and Cy5, respectively. Table S1). In the absence of the target, the two gating strands were spatially separated, the respectively labeled donor and acceptor fluorophores were separated, FRET was in the “off” state. Upon recognition of the target, the two gating strands were from open to closed state, bringing the dual fluorophores into close proximity and leading to high FRET efficiency.

Scheme 1. Construction (A) and Schematic Illustration (B) of DNA TP Nanoprobe for ATP Sensing in Living Cells

Preparation and Characterization of DNA TP Nanoprobe. To form the DNA TP nanoprobe, equimolar amounts of the three 96-base clips sequences (S1, S2, S3, Table S1) were combined and annealed at a temperature range from 95 °C to 4 °C over 3 h. Assembled structures were characterized by native PAGE (Figure S1A). Then the two functional oligonucleotides single-stranded strands (Apt1, Apt2. Table S1) were loaded onto the TP by a annealing at a temperature range from 95 °C to 4 °C

ACS Paragon Plus Environment

Analytical Chemistry

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

over 4 h. The size of the rigid TP construction was estimated to be 10.4 nm if the length of the 20-base TP edge was presumably 6.8 nm (Figure S3). Dynamic light scattering (DLS) of the DNA TP nanostructure shows a near monodisperse particle population with a hydrodynamic radius of (16.0 ± 2.0 nm) (Figure S4), and atomic force microscopy (AFM) shows collapsed circular objects with height values of (1.3 ± 0.3 nm) (Figure 2A and 2B). Figure 1 demonstrates the stepwise assembly of two capture sequences and a split aptamer on the TP, with corresponding decreases in gel mobility from lane 1 to lane 5. The results illustrated that our DNA TP nanoprobe had been successfully assembled by observing lane 5 TP-Apt1/Apt2 (363 nt) mobility corresponded to the marker position about 200 bp. Meanwhile, the another gel was run in the absence and in the presence, respectively, of 5 mM ATP, which demonstrated that DNA TP nanoprobe have good target-binding ability (Figure S1B). And its response to ATP was investigated in the further fluorescence experiment.

Figure 1. Analysis by 5% native PAGE. Line 0: 20bp DNA ladder; Line 1: S1; Line 2: S1+S2; Line 3: TP; Line 4: TP+Apt1; Line 5: TP+Apt1+Apt2.

ACS Paragon Plus Environment

Page 10 of 22

Page 11 of 22

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

Figure 2. (A) Characterization of DNA TP Nanoprobe by atomic force microscopy (AFM); Scale bars: 100 nm. (B) Line profile of a cross section in the AFM imaging in panel B.

In Vitro Fluorescence Response of DNA TP Nanoprobe. The DNA TP nanoprobe was used for the in vitro detection of ATP prior to sensing in living cells. At first, as shown in Figure 3A, the FRET signals were recorded according to the concentration of ATP in the range of 0.03-5 mM. In the absence of ATP, the DNA TP nanoprobe showed very low FRET signals. Upon recognition of ATP, the fluorescence emission intensity of Cy3 (donor) at 566 nm weakened rapidly, whereas the fluorescence emission intensity of Cy5 (acceptor) at 667 nm enhanced gradually. Figure 3B demonstrates the relationship between fluorescence emission ratio of acceptor to donor (FA/FD) and different concentrations of ATP. An almost 5-fold increase in FA/FD signal (from 0.09 to 0.48) was observed when the concentration of ATP increased from 0.03 mM to 5 mM. Meanwhile, the fluorescence emission ratio was found to be linear with the concentration of ATP in the range of 0.03-2 mM, and the limit of detection (LOD) was calculated to be 0.03 mM (R2 = 0.994). The concentration of ATP in living cells is typically 1-10 mM (most commonly between 2–3 mM).46-48 As listed in Table S2, which indicated the concentration of ATP in Hela cells is less than 3 mM. Therefore, the linear range of the DNA TP Nanoprobe ideally met the need for the sensing of ATP in living cells. Since it is necessary to study the response speed and selectivity between the DNA TP nanoprobe and the target. Real-time recording of the fluorescence emission

ACS Paragon Plus Environment

Analytical Chemistry

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

intensity changes of DNA TP nanoprobe as a function of time upon addition of ATP (5 mM) was carried out, the excitation wavelength was fixed at 525 nm, and the emission wavelength was 667 nm (Figure S5). Results showed the function between ATP and the DNA TP nanoprobe was finished within 10 min, suggesting that the DNA TP nanoprobe can response to the target rapidly. Meanwhile, control experiments revealed that analogue molecules of ATP, for example, cytidine triphosphate (CTP), guanosine triphosphate (GTP), and uridine triphosphate (UTP), only led to weak responses, suggesting the high selectivity of this DNA TP nanoprobe (Figure S6). In short, the DNA TP nanoprobe has the potential for further application in detecting ATP in living cells.

Figure 3. (A) Fluorescence emission spectra of 50 nM DNA TP nanoprobes in the presence of different concentrations of ATP (0, 0.5, 1, 1.25, 1.5, 1.75, 2, 2.5, 3, 4, and 5 mM). (B) The relationship between fluorescence emission ratio of acceptor to donor (FA/FD) and different concentrations of ATP. The inset shows a good linear correlation (R2 = 0.994) from 0.03 to 2 mM.

Stability and Cell Viability Assay of the DNA TP Nanoprobes. Nuclease resistance of the nanoprobes is quite important for an imaging application in complex biological systems. To confirm the stability of the DNA TP nanoprobes, DNase I was used to incubate with DNA TP nanoprobes. DNase I is an important endonuclease that effectively degrades single and double stranded DNA molecules into small fragments by cleaving the phosphodiester bonds in the DNA backbone.49,50 According to

ACS Paragon Plus Environment

Page 12 of 22

Page 13 of 22

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

reported serum levels of DNase I was (0.36±0.20) U/mL,51 we respectively chosed a near or far higher than that of plasma concentration of DNase I. So the DNA TP nanoprobes were respectively treated with low concentration of DNase I (0.25 U/mL)and high concentration of DNase I (2.5 U/mL) at 37 °C with different time periods. As shown in Figure 4A, the bands remained clearly within 1 h incubation with 0.25 U/mL DNase I, which is accordance with previous studies that DNA TP is strongly resistant to degradation of endonucleases.30,39 In other words, the resistance contributes to the steric hindrance of the DNA triangular prism nanostructure leading to declined binding of enzymes to DNA. However, when the nanoprobes were treated with 2.5 U/mL DNase I, the bands were obviously weakening and was almost completely disappeared at around 40 min. In order to further illustrate the DNA TP nanoprobe has a better ability of anti-nuclease degradation, a control experiment with the non-associated ssDNA probe was given. As shown in Figure S7, the DNA TP nanoprobes and ssDNA probes were respectively treated with DNase I (0.25 U/mL), compared with the latter’s bands disappeared completely in 10 minutesthe, the former’s bands remained clearly. As shown in Figure 4B, 0-6 h FBS (10% v/v) degradation assay illustrated that the DNA TP nanoprobe is more stable than ssDNA probe. These results illustrate the enhanced ability of the 3D-DNA nanostructure to protect oligonucleotides against nuclease degradation. Meanwhile, toxicity is always a major factor for nanoprobe designed to image and monitor markers into biological experiments. Hence, the standard colorimetric MTS assay was employed for the evaluation of the cytotoxicity of DNA TP nanoprobe on HeLa cells. The absorbance of MTS at 490 nm relies on the activation degree of cells, and the cell viability of the control group was about 100%. As shown in Figure S8, Hela cells were incubated with different concentrations of the DNA TP nanoprobes (0, 50, 100, 200, 400 nM) for 24 h at 37 °C. Cell survival remained more than 90% after exposure to DNA TP nanoprobes of 4-fold higher concentrations than that of the nanoprobes used in the following experiments. Therefore, the prismatic scaffold provides an effective delivery vehicle for biomaterials transporting and

ACS Paragon Plus Environment

Analytical Chemistry

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

intracellular sensing.

Figure 4. PAGE analysis of (A) the low concentration of DNase I (0.25 U/mL) and high concentration of DNase I (5 U/mL) degradation assay products for DNA TP nanoprobe. (B) FBS (10% v/v) degradation assay products for DNA TP nanoprobe and ssDNA at 37 °C.

In Situ cell Imaging of the DNA TP Nanoprobe. After established the signaling capability of the scaffolded DNA triangular prism, we further studied whether such nanostructures could function in cells for intracellular detection. We designed a functional nanomechanical DNA triangular prism device that encapsulating two split aptamers as binding site for the response of ATP in living cells (utilizing Hela cells as a model). In this case, the cellular uptake ability of the DNA TP nanoprobe was investigated. Both 50 nM of DNA TP nanoprobe and 50 nM of nude split aptamers were incubated with Hela cells for 4 h at 37 °C. As shown in Figure 5, cells treated with nude split aptamers displayed a very weak Cy3 fluorescence signal (green) and almost no Cy5 fluorescence signal (red), this illustrated that the nude ssDNA probe was not suitable for imaging intracellular ATP, because of its poor permeability and low biostability. In contrast, cells treated with DNA TP nanoprobe displayed obvious Cy3 fluorescence signal (green) and Cy5 fluorescence signal (red), indicating that intracellular ATP brought the two dyes close together, enabling efficient FRET from

ACS Paragon Plus Environment

Page 14 of 22

Page 15 of 22

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

Cy3 to Cy5. Such ATP-induced increase of Cy5 fluorescence and decrease of Cy3 fluorescence was in accordance with the in vitro experiments in 1×TAE/Mg2+ solution. As ATP is the major energy currency molecule in living cells, to further confirm the fluorescence signal is due to the endogenously produced ATP of the Hela cells, an assay for the DNA TP nanoprobe to detect the changes in ATP levels is essential. In the following experiments, an in situ ATP imaging was designed. It has been reported that oligomycin is a well-known inhibitor of ATP and that Ca2+ is a commonly used ATP inducer.52,53 Hence, before incubated with 50 nM DNA TP nanoprobe, Hela cells were treated with 10 µM oligomycin or with 5 mM Ca2+ for 30 min at 37 °C. As shown in Figure 6, the higher FRET signal in the Ca2+-treated cells (bottom) and lower FRET signal in the oligomycin-treated cells (middle) could be obviously observed compared with those in the untreated cells (top). These results have illustrated the DNA TP could deliver the aptamer probe into Hela cells and successfully achieve sensing of ATP in living cells.

Figure 5. Fluorescence images of Hela cells after incubation with 50 nM DNA TP nanoprobe (top), and 50 nM ATP-Apt1/Apt2 (bottom) for 4 h at 37 °C. Scale bar: 10 µm.

ACS Paragon Plus Environment

Analytical Chemistry

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 6. Fluorescence images of Hela cells treated with medium (top), 10 µM oligomycin (middle), and 5 mM Ca2+ (bottom), followed by incubation with 50 nM DNA TP nanocomplex for 4 h at 37 °C. Scale bar: 10 µm.

Hence, the results above indicated that the DNA TP nanoprobe is superior to the traditional strategy of molecular beacon-based biosensor construction. Firstly, the DNA TP nanoprobe exhibited a low detection limit and high selectivity toward ATP in the vitro assay. Secondly, in situ cell imaging indicated that the DNA TP nanoprobe exhibited high cellular permeability, fast response, and successfully realized “FRET off” to “FRET on” sensing of ATP in living cells owing to the strong protecting capability to nucleic acids from enzymatic cleavage and the excellent biocompatibility of the TP. In contrast, the nude ssDNA probe showed low cellular permeability and could not detect FRET signal. Finally, but arguably most importantly, the DNA TP nanoprobe constructed by organic combination of the excellent biological properties of DNA nanostructures, the low background split aptamers, and the FRET “off” to “on” sensing mechanism, revealed unparalleled advantages. Compared with the traditional approaches incorporating inorganic nanomaterials, such as gold nanoparticles and

graphene, the complex steps for the preparation and

functionalization of these nanomaterials could be avoid, the stability of nucleic acids were enhanced, and false-positive signals could be basically avoided. Overall, the

ACS Paragon Plus Environment

Page 16 of 22

Page 17 of 22

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

methods and the design presented here provide an overview of the steps involved in functional DNA-minimal cage design, while providing an adaptable nanostructure for potential biological applications.

CONCLUSION In summary, we have developed a DNA triangular prism encapsulating a split aptamer for adenosine triphosphate (ATP) sensing in living cells. In vitro assays demonstrated that the DNA TP nanoprobe was a stable, sensitive, and selective biosensor for quantitative detection of ATP, and the DNA TP is able to protect its cargo against site-specific cleavage and nuclease degradation. Moreover, the confocal fluorescence microscopy experiments with Hela cells further indicated that the DNA TP nanoprobe was efficiently delivered into living cells and worked as an in situ FRET “off” to “on” biosensor for specific, high-contrast imaging of target molecules. We foresee these 3D DNA nanoprobes will become a highly generic and versatile structure for further development of DNA nanotechnology and DNA computers that function both in vitro and in vivo.

ACKNOWLEDGMENTS

This research was supported by the National Natural Science Foundation of China (No.21675043), the Foundation for Innovative Research Groups of NSFC (Grant 21521063) and the science and technology project of Hunan Province (2016RS2009, 2016WK2002).

ASSOCIATED CONTENT The Supporting Information is available free of charge via the Internet at http://pubs.acs.org.

ACS Paragon Plus Environment

Analytical Chemistry

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

Supporting Information: DNA synthesis, construction of the TP nanostructure, DNA sequences, gel electrophoresis, and assembly schematic; DLS, fluorescence spectra, and cell viability assay.

REFERENCES (1) Feng, X.; Jia, Y.; Cai, P.; Fei, J. B.; Li, J. B. Acs Nano 2016, 10, 556-561. (2) Palleros, D. R.; Raid, K. L.; Shi, L.; Welch, W. J.; Fink, A. L. Nature 1993, 365, 664−666. (3) Desai, A.; Verma, S.; Mitchison, T. J.; Walczak, C. E. Cell 1999, 96, 69−78. (4) Van Wylen, D. G.; Park, T. S.; Rubio, R.; Berne, R. M. J. Cereb. Blood Flow Metab. 1986, 6, 522-528. (5) Gourine, A. V.; Llaudet, E.; Dale, N.; Spyer, K. M. Nature 2005, 436, 108–111. (6) Zhu, C.; Zhao, Y.; Yan, M. M., Huang, Y. F.; Yan, J.; Bai, W. H.; Chen, A. L. Anal. Bioanal.

Chem. 2016, 408, 4151-4158. (7) Bratu, D. P.; Cha, B. J.; Mhlanga, M. M.; Kramer, F. R.; Tyagi, S. Proc. Natl. Acad. Sci. USA

2003, 100, 13308−13313. (8) Medley, C. D.; Drake, T. J.; Tomasini, J. M.; Rogers, R. J.; Tan, W. H. Anal. Chem. 2005, 77, 4713−4718. (9) Bock, L. C.; Griffin, L. C.; Latham, J. A.; Vermaas, E. H.; Toole, J. J. Nature 1992, 355, 564– 566. (10) Chen, C. H. B.; Chernis, G. A.; Hoang, V. Q.; Ralf, L. Proc. Natl. Acad. Sci. USA 2003, 100, 9226–9231. (11) Shangguang, D.; Li, Y.; Tang, Z.; Cao, Z.; Chen, H.; Mallikaratchy, P.; Sefah, K.; Yang, C. J.; Tan, W. Proc. Natl. Acad. Sci. USA 2006, 103, 11838–11843. (12) Zhang, H.; Ma, Y.; Xie, Y.; An, Y.; Huang, Y.; Zhu, Z.; Yang, C. J. Sci. Rep. 2015, 5, 1−5.

ACS Paragon Plus Environment

Page 18 of 22

Page 19 of 22

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

(13) Wang, Y.; Wu, Z.; Liu, S.; Chu, X. Anal. Chem. 2015, 87, 6470−6474. (14) Lei, Y. L.; Tang, J. L.; Shi, H.; Ye, X. S.; He, X. X.; Xu, F. Z.; Yan, L. A.; Qiao, Z. Z.; Wang, K. M. Anal. Chem. 2016, 88, 11699-11706. (15) Stojanovic, M. N.; Prada, P. D.; Landry, D. W. J. Am. Chem. Soc. 2000, 122, 11547-11548. (16) Zuo, X.; Xiao, Y.; Plaxco, K.W. J. Am. Chem. Soc. 2009, 131, 6944–6945. (17) Lin, Z.; Luo, F.; Liu, Q.; Chen, L.; Qiu, B.; Cai, Z.; Chen, G. Chem. Commun. 2011, 47, 8064–8066. (18) Du, Y.; Guo, S.; Qin, H.; Dong, S.; Wang, E. Chem. Commun. 2012, 48, 799–801. (19) Zhu, Z.; Ravelet, C.; Perrier, S.; Guieu, V. R.; Fiore, E.; Peyrin, E. Anal. Chem. 2012, 84, 7203–7211. (20) Sato, Y.; Zhang, Y.; Nishizawa, S.; Seino, T.; Nakamura, K.; Li, M.; Teramae, N. Chem. Eur.

J. 2012, 18, 12719–12724. (21) Sharma, A. K.; Kent, A. D.; Heemstra, J. M. Anal. Chem. 2012, 84, 6104–6109. (22) Zhang, J.; Wang, L. H.; Pan, D.; Song, S. P.; Boey, F. Y. C.; Zhang, H.; Fang, C. Small 2008,

4, 1196–1200. (23) Sharma, A. K.; Heemstra, J. M. J. Am. Chem. Soc. 2011, 133, 12426–12429. (24) Zhang, H.; Ma, Y.; Xie, Y.; An, Y.; Huang, Y.; Zhu, Z.; Yang, C. Sci. Rep. 2015, 5, 1−5. (25) Li, J.; Fan, C.; Pei, H.; Shi, J.; Huang, Q. Adv. Mater. 2013, 25, 4386−4396. (26) Wu, X.; Chen, J.; Wu, M.; Zhao, X. Theranostics 2015, 5, 322−344. (27) Yin, P.; Hariadi, R. F.; Sahu, S.; Choi, H. M. T.; Park, S. H.; LaBean, T. H.; Reif, J. H.

Science 2008, 321, 824-826. (28) Liang, L.; Li, J.; Li, Q.; Huang, Q.; Shi, J. Y.; Yan, H.; Fan, C. H. Angew. Chem., Int. Ed.

2014, 53, 7745-7750.

ACS Paragon Plus Environment

Analytical Chemistry

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

(29) Hamblin, G. D.; Rahbani, J. F.; Sleiman, H. F. Nat. Commun. 2015, 6, 7065-7072. (30) Bujold, K. E.; Hsu, J. C.; Sleiman, H. F. J. Am. Chem. Soc. 2016, 138, 14030-14038. (31) Meng, H. M.; Liu, H.; Kuai, H. L.; Peng R. Z.; Mo, L. T.; Zhang, X. B. Chem. Soc. Rev. 2016,

45, 2583-2602. (32) Wei, B.; Dai, M.; Yin, P. Nature 2012, 485, 623-626. (33) Winfree, E.; Liu, F.; Wenzler, L. A.; Seeman, N. C. Nature 1998, 394, 539-544. (34) Edwardson, T. G. W.; Carneiro, K. M. M.; Mclaughlin, C. K.; Serpell, C. J.; Sleiman, H. F.

Nat. Chem. 2013, 5, 868-875. (35) Nishikawa, M.; Rattanakiat, S.; Takakura, Y. Adv. Drug Delivery Rev. 2010, 62, 626–632. (36) Pei, H.; Liang, L.; Yao, G. B.; Li, J.; Huang, Q.; Fan, C. H. Angew. Chem., Int. Ed. 2012, 51, 9020-9024. (37) Edwardson, T. G. W.; Kai, L. L.; Bousmail, D.; Serpell, C. J.; Sleiman, H. F. Nat. Chem. 2016,

8, 162-170. (38) McLaughlin, C. K.; Hamblin, G. D.; Sleiman, H. F. Chem. Soc. Rev. 2011, 40, 5647-5656. (39) Conway, J. W.; Mclaughlin, C. K.; Castor, K. J.; Sleiman, H. F. Chem. Commun. 2013, 49, 1172-1174. (40) Liu, Y.; Chen, Q. S.; Liu, J. B.; Yang, X. H.; Guo, Q. P.; Li, L.; Liu, W.; Wang, K. M. Anal.

Chem. 2017, 89, 3590−3596. (41) He, L.; Lu, D. Q.; Liang, H.; Xie, S. T.; Luo, C.; Hu, M. M.; Xu, L. J.; Zhang, X. B.; Tan, W. H. ACS Nano 2017, 11, 4060-4066. (42) Dave, N.; Liu, J. Chem. Commun. 2012, 48, 3718-3720. (43) Zhang, Z.; Liu, J. Acs Appl. Mater. Interfaces 2016, 8, 6371-6378. (44) Zhang, Z. J.; Oni, O.; Liu, Nucleic Acids Res. 2017, 45, 7593-7601.

ACS Paragon Plus Environment

Page 20 of 22

Page 21 of 22

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

(45) Peng, R. Z.; Wang, H. J.; Lyu, Y. F.; Xu, L. J.; Liu, H.; Kuai, H. L.; Liu, Q. L.; Tan, W. H. J.

Am. Chem. Soc. 2017, DOI: 10.1021/jacs.7b07485. (46) Imamura, H.; Huynh Nhat, K. P.; Togawa, H.; Saito, K.; Iino, R.; Kato-Yamada, Y.; Nagai, T.; Noji, H. Proc. Natl. Acad. Sci. USA 2009, 106, 15651−15656. (47) Qiang, W. B.; Hu, H. T.; Sun, L.; Li, H.; Xu, D. K. Anal. Chem. 2015, 87, 12190-12196. (48) Traut, T. W. Mol. Cell. Biochem. 1994, 140, 1-22. (49) Keum, J. W.; Bermudez, H. Chem. Commun. 2009, 45, 7036−7038. (50) Li,N.; Wang, M.; Gao, X.; Yu, Z.; Pan, W.; Wang, H.; Tang, B. Anal. Chem. 2017, 89, 6670-6677. (51) Cherepanova, A.; Tamkovich, S.; Pyshnyi, D.; Kharkova, M.; Vlassov, V.; Laktionov, P. J.

Immunol. Methods 2007, 325, 96–103. (52) Gong, Y. X.; Sohn, H.; Xue, L.; Firestone, G. L.; Bjeldanes, L. F. Cancer Res. 2006, 66, 4880−4887. (53) Ainscow, E. K.; Rutter, G. A. Diabetes 2002, 51, S162−S170.

ACS Paragon Plus Environment

Analytical Chemistry

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

For TOC only

ACS Paragon Plus Environment

Page 22 of 22