Single Molecule, Real-Time Dissecting of Peptide Nucleic Acids-DNA

Subscriber access provided by UNIV OF NEW ENGLAND ARMIDALE

Article

Single Molecule, Real-Time Dissecting of Peptide Nucleic Acids-DNA Duplexes with a Protein Nanopore Tweezer Andrei Ciuca, Alina Asandei, Irina Schiopu, Aurelia Apetrei, Loredana Mereuta, Chang Ho Seo, Yoonkyung Park, and Tudor Luchian Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b01568 • Publication Date (Web): 25 May 2018 Downloaded from http://pubs.acs.org on May 25, 2018

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

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

Page 1 of 10 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

Single Molecule, Real-Time Dissecting of Peptide Nucleic Acids-DNA Duplexes with a Protein Nanopore Tweezer Andrei Ciuca1,#, Alina Asandei2#, Irina Schiopu2#, Aurelia Apetrei1, Loredana Mereuta1, Chang Ho Seo3, Yoonkyung Park4,* & Tudor Luchian1,* 1

Department of Physics, Alexandru I. Cuza University, Iasi, Romania. Interdisciplinary Research Department, Alexandru I. Cuza University, Iasi, Romania. 3 Department of Bioinformatics, Kongju National University, Kongju, South Korea. 4 Department of Department of Biomedical Science and Research Center for Proteinaceous Materials (RCPM), Chosun University, Gwangju, Korea. 2

ABSTRACT: Peptide nucleic acids (PNAs) are artificial, oligonucleotides analogues, where the sugar-phosphate backbone has been substituted with a peptide-like N-(2- aminoethyl)glycine backbone. Due to their inherent benefits, such as increased stability and enhanced binding affinity towards DNA or RNA substrates, PNAs are intensively studied and considered beneficial for the fields of materials and nanotechnology science. Herein, we designed cationic polypeptide-functionalized, 10-mer PNAs, and demonstrated the feasible detection of hybridization with short, complementary DNA substrates, following analytes interaction with the vestibule entry of an α-hemolysin (α-HL) nanopore. The opposite charged state at the polypeptide-functionalized PNA-DNA duplex extremities, facilitated unzipping of a captured duplex at the lumen entry of a voltage-biased nanopore, followed by monomers threading. These processes were resolvable and identifiable in real-time, from the temporal profile of the ionic current through a nanopore accompanying conformational changes of a single PNA-DNA duplex inside the α-HL nanopore. By employing a kinetic description within the discrete Markov chains theory, we proposed a minimalist kinetic model to successfully describe the electric force-induced strand separation in the duplex. The distinct interactions of the duplex at either end of the nanopore present powerful opportunities for introducing new generations of force-spectroscopy nanopore-based platforms, enabling from the same experiment duplex detection and assessment of interstrand base pairing energy.

A long-sought goal of biotechnology and nanotechnology is to deliver label-free, single-molecule screening protocols enabling sensing, discriminating, and manipulating single molecules in aqueous solutions. To meet this challenge, one of the richest approaches makes use of nanopores, either protein1–5 or solid-state based.6,7 The working principle of the approach generally follows this sequence of events: (i) an electric field drives a macromolecule of interest towards the nanopore (ii) the transient confinement of the macromolecule inside the nanopore’s inner volume leads to the displacement of a corresponding volume of solvent, entailing alterations of the electrical resistance of the nanopore (iii) under voltage-clamp conditions across the nanopore, reversible changes in the ionic electrical current measured across the nanopore occur, seen as ionic current blockade events. The volumetric analysis of the amplitude of such blockade events, and the statistical analysis of blockade durations and blockade-events frequency, made usually within the Markov state models,8–10 provides knowledge about the physical and chemical features of the studied macromolecule, with high temporal (microsecond scale) and spatial (nanometer scale) resolution.

The α-hemolysin (α-HL) protein secreted by Staphylococcus aureus11 oligomerizes and self-assembles in target membrane to produces a water-filled mushroom-like heptameric pore. The knowledge of α-HL’s crystal structure, its intrinsic robustness and physico-chemical properties, transformed the αHL into a powerful tool for probing single-molecule biophysics phenomena at nanoscale.2–4,12–19 This undertaking has been further facilitated by the possibility to control the speed and the direction of motion of single molecules accompanying their detection with the α-HL nanopore.20–22 With particular relevance to the growing field of nucleic acids nanotechnology, peptide nucleic acids (PNA), which are structural DNA analogues containing an uncharged N-(2aminoethyl)- glycine-based pseudopeptide backbone, have been shown to mimic DNA in forming Watson-Crick complementary duplexes with normal DNA.23 PNAs demonstrated tremendous potential for use in molecular diagnostics and antisense therapeutics24 due to its greater binding affinity, selectivity, as well as its resistance to degradation by nucleases and proteases.25,26 The process of hybridization, in which an oligonucleotide probe recognizes and binds to its complemen-

1 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

tary target, has been successfully probed using nanopores,27,28 the mechanisms for the trapping, unzipping, and translocation of nucleic acids in the nanopore were probed29 and the particular PNA-based hybridization was shown to be particularly useful for selective detection of dsDNA30 or miRNA in the nucleic acid mixture.31 To turn PNAs into powerful tools for building novel supramolecular structures, facile and cheap ways to detect their hybridization and measure the binding energy to different substrates, are paramount. To this end, the very principles outlined for molecule detection with nanopores, were proposed as a viable alternative in the realm of single-molecule force spectroscopy. An electric field is used to drive a charged molecule through a nanopore, forces are then exerted on a nanopore-confined molecule that can lead to its unfolding and subsequent breakage. The response of the system can be followed as a function of time, to extract kinetic information which later can be used to infer information regarding the molecules rupture energy.32,33 This technique was successfully employed to study the unfolding of DNA hairpins and DNA duplexes,34 DNA–ligand complexes,35,36 it utilized duplex unzipping to detect metal ion binding,37 and base-pair mismatches.38 In particular, duplex unzipping through the αHL has attracted much interest, and the unzipping kinetics and base pairing energy of duplex DNA have been extensively explored.39–43

Page 2 of 10

Buffer solutions. DNA and PNA samples were dissolved in a 1 M NaCl solution prepared in ultra-pure water, buffered with TE (10 mM Tris, 1 mM EDTA) at a pH=7.9. For the electrophysiology experiments, a 3 M KCl electrolyte solution was prepared in ultra-pure water, buffered with 10 mM HEPES to a pH =7.4. Sample preparation. DNA1 and DNA3 oligonucleotide samples in dried form, were resuspended in 1 M NaCl/TE buffer to obtain stock solutions of 200 µM and 100 µM, respectively. Stock solutions of PNA3 were obtained by hydrating the dry samples in the same buffer up to a concentration of 200 µM. All liquid samples were vigorously stirred using a Stuart vortex-mixer with BioCote (Sigma–Aldrich, Germany) at 1400 rpm for 3 minutes each, then heated up to 95 °C for 20 min to improve rehydration. Aliquots of solution to be used in electrophysiology experiments were transferred into new vials and the remaining stock solutions were stored at -20 °C until further use. Prior to use in electrophysiology experiments, aliquots were heated to 95 °C using an IKA Digital Block Heater (Cole-Parmer, US) and slowly cooled down to 22 °C. In addition, the PNA3-DNA1 hybridization was measured in a similar buffer as used in electrophysiology. Before hybridization, all molecules were separately annealed by rapidly heating each sequence to 95o C and slowly cooling to 22o C, using an IKA Digital Block Heater (Cole-Parmer, US). Hybridization was then observed by monitoring the decrease in UV absorbance at 260 nm upon mixing distinct molar concentrations of PNA3 and DNA1 solutions, with Thermo Scientific, NanoDrop OneC spectrophotometer (data not shown).

In this report, we used polycationic peptide-functionalized PNAs, and utilized the α-HL nanopore as a single-molecule sensor, to detect and investigate the voltage-dependent unzipping of the oppositely charged PNA-DNA duplex. We reported previously on a similar approach which we dubbed ‘nanopore tweezing’, and demonstrated that by placing opposite electric charges at a peptide ends, leading to oppositelyoriented forces acting on the analyte placed in an electric field, a dramatic slow-down of its motion across the α-HL nanopore ensues.44,45 We thus reasoned that by using a similar strategy, not only a dipolar molecular duplex would be slowed-down, but also more efficiently dissociated. We demonstrate herein that electric forces from the applied transmembrane potential, acting oppositely at both ends of a hybridized PNA-DNA duplex trapped inside the α-HL nanopore, facilitate unzipping. From the analysis of the current blockage level changes accompanying the unzipping of a single PNA-DNA duplex inside the nanopore's β-barrel, and the time it takes to unzip, we proposed the mechanism, kinetic scheme and parameters that could describe strand separation.

Table 1. Primary sequences of the polypeptide-functionalized PNA and ssDNA strands. The red-marked region within the DNA1 represents the complementary domain to the PNA3. Within same region of DNA3, we underlined the mismatches introduced to the complementary domain. The poly-C tails were introduced at the 5’ end in both DNA1 and DNA3 sequences, to provide a free overhang sequence on the duplexes, oppositely charged with respect to the poly-Arg tail attached to the PNA3. Polynucleotide

Primary sequence

Mw (g/mol)

PNA3

Ac-(R)9–5’-GTGATATACG-3’

4214.4

EXPERIMENTAL SECTION

DNA1

5’-CCCCCCCCGTATATCAC-3’

5014

Materials. We designed the polypeptide-functionalized PNA and DNA sequences, which were subsequently synthesized and purified by Panagene Inc., Republic of Korea (PNAs) and Sigma-Aldrich, Germany (DNAs). The primary sequences and molecular weights are shown in Table 1. The 1,2-diphytanoylsn-glycerophosphocholine (DPhPC) lipid was obtained from Avanti Polar Lipids, Alabaster, AL, USA and the α-hemolysin (α-HL), potassium chloride (KCl), sodium chloride (NaCl), ultra-pure water (DNAase and RNAase free), n-pentane, hexadecane, EDTA, and buffers (Tris and HEPES) were purchased from Sigma–Aldrich, Germany.

DNA3

5’-CCCCCCCGTGTTTTTCG-3’

5067

Single-molecule electrophysiology. The planar lipid membranes were formed by using the Montal-Mueller technique.44,46 Both chambers of the bilayer cell were filled with equal volumes of the 3 M KCl electrolyte solution. α-HL (~ 0.5 – 2 µL) was added to the grounded, cis-chamber from a monomeric stock solution made in 0.5 M KCl. After the successful insertion of a single heptameric α-HL nanopore into the lipid membrane, ssDNAs and PNAs were added from the stock solution to either the cis- or trans-side of the membrane.


Page 3 of 10 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 nanopore was subjected to distinct potential difference values, and the corresponding ionic currents were recorded via two Ag/AgCl electrodes connected to an Axopatch 200B (Molecular Devices, USA) amplifier, in voltage-clamp mode. When analytes were added to the cis-chamber, the membrane was clamped to negative voltage values ranging from ∆V = 100 mV to ∆V = - 200 mV. In this protocol, we first added PNA3 (3 µM) to the cis solution and ionic currents through the α-HL nanopore were recorded at various potentials. We then added either DNA1 or DNA3 to the cis solution at a concentration of 9 µM, thus establishing a PNA:DNA ratio of 1:3, and carried out ionic current recordings as above. When analytes were added to the trans side of the membrane, positive voltages were applied. In control experiments, current fluctuations induced by 3 µM PNA3 – nanopore interactions were recorded at positive potential differences across the membrane, ranging from ∆V = + 100 mV to ∆V = + 200 mV. DNA1 was then added incrementally to the trans chamber to achieve final PNA3:DNA1 molar ratios of 1:3, 1:5 and 1:10, respectively.46 During all experiments involving PNA-DNA mixtures, the molecules were allowed to interact for ~15 min, to reach the equilibrium of the hybridization reaction before starting the recordings. Addition of pre-hybridized PNA3DNA1 duplexes formed as described above (Sample preparation) to the trans solution, resulted in largely similar events when interacting with the nanopore (data not shown). Electrical signals were low-pass filtered at 10 kHz and digitized with a NI PCI 6221, 16-bit acquisition board (National Instruments, USA) at a sampling frequency of 50 kHz, with a virtual instrument developed within the LabVIEW 8.20 (National Instruments, USA). To reduce the contributions of the electrical and mechanical noise, the recording system was shielded in a Faraday cage (Warner Instruments, USA), and placed on a vibration-free platform (Bench Mate 2210, Warner Instruments, USA).

Figure 1. Uni-molecular view of interactions between cis added PNA3 molecules and the α-HL. (a) The electric field across the negatively-biased nanopore acts on the positively charged poly-arginine chain from the PNA3, and the resulting force drives the molecule inside the nanopore’s vestibule. (b) Selected trace illustrating the reversible PNA3 – α-HL interactions, showed as upwardly oriented current spikes, recorded at ∆V = - 160 mV, upon cis addition of PNA3 (3 µM). (c) The all-points histogram reflecting different conductive states of α-HL measured in the presence of the cis-added PNA3, namely: free nanopore (O), the presence of a PNA3 inside the nanopore’s vestibule (V) and two blockades states (B1 and B2) associated with the presence of the PNA3 in the constriction region on the nanopore during passage. (d) The vestibule-lumen transitions (VL) (left-hand side) are indicative of steps during which a PNA3 molecule passes through the nanopore, upon capture inside the vestibule. The vestibulelumen-vestibule transitions (VLV) (middle) show the sojourn of the PNA3 molecule which, upon capture inside the nanopore’s vestibule and partial threading across the nanopore’s constriction region, exits the nanopore through the cis side. The right-hand side trace depicts events whereby the PNA3 molecules briefly enter the nanopore’s vestibule and returns to the cis side, without reaching the constriction region of α-HL.

(e) Voltage dependence of the average dwell times ( ) associated to the vestibule-lumen transitions (VL) of a PNA3 molecule. (f) Voltage dependence of the average dwell time

reflecting vestibule-lumen-vestibule transitions (VLV) ( ). Upper and lower confidence limits mark the 95 % confidence intervals for the estimated average dwell times.

RESULTS AND DISCUSSION Interaction of the PNA-DNA duplex at the α-HL’s vestibule allows hybridization detection, while the α-HL’s lumen facilitates duplex unzipping We show in Fig. 1, panel a, the experimental principle of PNA–DNA duplex detection using the α-HL nanopore. In a first series of experiments, the single-stranded cationic polypeptide-functionalized PNA (called herein PNA3, see Table 1) was presented on the grounded, cis side of the nanopore. The PNA3 molecules are electrophoretically driven towards the vestibule entrance of the negatively biased nanopore, by the transmembrane potential.

The PNA3-nanopore interactions were probed through the reversible reduction in the ionic current through the nanopore.



Closer examination of these events shows multiple types of blockade levels, as they are discussed in Fig. 1. When the complementary ssDNA sequence (called herein DNA1, see Table 1) was added in cis solution, the signature of PNA3induced blockade events through the nanopore changed (Fig. 2, panels a-c).

Page 4 of 10

For the purpose of successfully capturing and unzipping PNA3-DNA1 complexes with the nanopore, all analytes were added to the trans side of the membrane, and positive transmembrane potentials applied across the nanopore. We reasoned that the PNA3-DNA1-nanopore interactions would occur distinctly, mainly due to the negatively charged entrance of the α-HL’s lumen at neutral pH.

Figure 2. The single-molecule, PNA3:DNA1 hybridization detection with the α-HL. Panels (a-c) illustrate that upon PNA3-DNA1 (1:3) duplex formation in cis solution, the ensuing decrease in the bulk concentration of free PNA3 leads to a corresponding decrease in the PNA3-α-HL reversible interactions as compared to control experiments (PNA3 alone, Fig.1, panels (b-c), at ∆V= - 160 mV. Panels (d-f) demonstrate that at a similar concentration ratio (PNA3-DNA3 (1:3)) and applied potential, the cis side addition of DNA3 leaves the occurrence of PNA3-α-HL interactions largely un-changed, when compared to control experiments (PNA3 alone, Fig.1, panels b-c).

Figure 3. Uni-molecular view of the trans-added PNA3 and PNA3-DNA1 interaction with the positively-biased α-HL. (a) Schematic representation of the trans-added PNA3 interacting with the α-HL under positively applied potentials. The association process is driven by the electrophoretic force ), and the electrostatic attractions ( ) between the ( positively charged poly-arginine chain on the PNA and the negatively charged trans opening of the α-HL (-7 |e-|, see text). (b) Selected recording showing reversible blockades induced by the interaction of trans-added PNA3 (3 µM) with the α-HL at ∆V = + 160 mV. (c) Cartoon representation of the PNA3DNA1 duplex interaction with a single α-HL. The cumulative effect of and overcomes the electrophoretic force acting oppositely on the negatively charged DNA1 ( ). (d) Selected current fluctuations recorded through the α-HL at ∆V = + 160 mV, upon trans-chamber addition of PNA3 (3 µM) and DNA1 (30 µM).

We attributed this to a depletion of free PNA3 molecules in bulk solution, followed hybridization with DNA1. The hybridized PNA3-DNA1 duplex carries a net negative charge, due to the larger negative charge on the DNA1 backbone (bare charge of - 17│e-│) as compared to the positive charge on the polypeptide-functionalized PNA1 (bare charge of + 9│e-│). In turn, this leads to the transport of the PNA3-DNA1 duplex away from the α-HL’s vestibule, along with the non hybridized DNA1, under the electrophoretic force ( ). Note that the electric signatures of the nanopore transiently blocked by the cis-added PNA3 in absence (Fig. 1, panel c) or DNA1 addition (Fig. 2, panel c), did not reveal major and reproducible differences in terms of relative extent of block or blockade duration. Further to this, addition of non-complementary ssDNA sequence (DNA3, see Table 1) in cis solution barely affects the electric signature of PNA3 detection (Fig. 2, panels d-f), suggesting that DNA3 does not diminishes through hybridization the bulk concentration of free PNA3. This observation suggests specificity of ssDNA detection via interaction with complementary polypeptide-functionalized PNA, required for real-time PNA–DNA hybridization detection. The negatively charged ssDNA molecules and the hybridized duplexes present in bulk are electrophoretically driven away from the nanopore’s entrance on the cis side, do not interfere with the PNA-nanopore interactions, thus imparting high sensitivity in detecting hybridization.

Fig. 3 illustrates the occurrence of PNA3 interactions with the α-HL’s β-barrel in control conditions (absence of the complementary DNA1 substrate) and presence of DNA1 in the trans solution. Counterintuitively, the frequency of events comprising the signature of blockades seen in the presence of the PNA3 alone (Fig. 3, panels a, b) compares very closely with that recorded upon the complementary DNA1 addition (Fig. 3, panels c, d). This is unexpected, since the bulk concentration the free PNA3 is expected to decrease upon hybridization with the complementary DNA1 substrate, and lead to a corresponding decrease in the number of blockade events (see also Fig. 2). To explain this, we posit that the blockade events recorded when both PNA3 and the complementary DNA1 strands were present in trans solution, reflect also the reversible interactions between the hybridized PNA3-DNA1 complexes and the nanopore. The physical basis of this is three-fold: (i) the PNA3-DNA1 duplex can orient itself in the electric field near the nanopore, and get funneled with the positively charged polypeptide from the PNA3 towards the nanopore’s lumen; (ii)


Page 5 of 10 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 electric field intensity in the bulk solution from the transmembrane potential is present within ~ 1 nm near the nanopore entrance, and scales sharply with the distance from the nanopore (r) as ~ r-2.47 Therefore, the PNA3-DNA1 duplex gets prone to associate to the nanopore, since the predominantly negatively charged DNA1 backbone of the complex - which situates farther from the nanopore’s entrance as compared to the PNA3 moiety - senses a diminished repelling force from the electric field near the nanopore, as compared to the attractive force acting on the PNA3; (iii) perhaps most importantly, the attractive electrostatic interactions ( ) between the polycationic peptide attached to PNA3, on the PNA3-DNA1 duplex, and the nanopore’s negatively charged lumen entrance (at neutral pH it carries a bare net charge of -7│e-│), facilitates the PNA3-DNA1 duplex association to the nanopore as depicted in Fig. 3, panel c. This is supported by previous results which demonstrated the possibility of controlling peptide capture and passage through the α-HL, by altering the electrostatic interactions between the protein’s mouth and either peptide’s terminus.48

sentative example of a PNA3-DNA1-induced blockade on the α-HL, showing a reversible lower-amplitude event (type ‘1’) or a more complex one consisting of a lower-amplitude blockade (type ‘2’) followed by a higher-amplitude blockade (type ‘3’). The type ‘3’ events have similar blockade amplitudes as those shown in panel c, and they were assigned to the ‘U’ sub-state in panel b. (e) Frequency of PNA3-DNA1induced blockade events (type ‘1’ and the type ‘2’ + type ‘3’ group in panel d) as a function of DNA1 concentration, at a fixed concentration of PNA3 (3 µM). The evidence supporting the accurate assignment of the blockade events shown in Fig. 4, panel c, as reflecting the PNA3nanopore interactions is two-fold. Firstly, the relative blockade amplitude such events seen when PNA3 and DNA1 were added together in buffer (denoted by ‘U’ in Fig. 4, panel b), was practically identical with that measured when PNA3 was added alone in the trans chamber (Figure 3, panel a) ( ∆IPNA3/IO = 0.995 ± 0.001, where IO represents the ionic current through an open nanopore – sub-state ‘O’, and ∆IPNA3=IOIU, with IU denoting the ionic current associated to the ‘U’ substate). This indicates a similar size of the molecules giving rise to the ionic current blockades exemplified in Fig. 3, panel a and Fig. 4, panel c. Secondly, the voltage-dependence of such blockade events, extracted from experiments carried out with either PNA3 alone, or PNA3 and DNA1 added together in buffer, is practically similar on both cases (Fig. S1, Supporting Information). This indicates a similar kinetic behavior for the molecular species blocking reversibly the nanopore as shown in Fig. 3, panel a and Fig. 4, panel c.

Interrogation of the PNA3-DNA1 duplex with the α-HL When the ionic current fluctuations shown in Fig. 3, panel d were examined more closely, two distinct signatures of current blockade were identified, which we assigned to the PNA3 alone and the PNA3-DNA1 duplex interacting with the α-HL (Fig. 4).

The second type of blockade events (Fig. 4, panel d) whose signature is more complex, was assigned to the hybridized PNA3-DNA1 duplex interacting with the nanopore. Such events were ascribed to a PNA3-DNA1 duplex partially entering the α-HL’s β-barrel with the PNA3-attached polypeptide tail head on, and then retraction back to the trans side (events denoted by ‘1’ in Fig. 4, panel d), or instances when following entrance to the nanopore (events denoted by ‘2’ in Fig. 4, panel d), unzipping of the PNA-DNA1 duplex occurs, facilitating the translocation of the unzipped PNA monomer across the nanopore (events denoted by ‘3’ in Fig. 4, panel d). A strong proof in favor of this hypothesis is that the frequency of blockade events shown in Fig. 4, panel d, increases with the augmentation of the DNA1 concentration relative to PNA3, and achieves a plateau around the 1:10 (PNA3-DNA1) molar ratio (Fig. 4, panel e). This verifies that such events reflect the nanopore interaction with the PNA3-DNA1 duplex, whose bulk concentration scales proportionally with the DNA1 concentration at a given PNA3 initial concentration. Note that the relative amplitude of PNA3-DNA1 duplex-induced blockade events (denoted by ‘1’ and ‘2’ in Fig. 4, panel d) are similar (∆IPNA3-DNA1;’1’/IO = 0.865 ± 0.018; ∆IPNA3-DNA1;’2’/IO = 0.868 ± 0.008, where ∆IPNA3-DNA1;’1’ = ∆IPNA3-DNA1;’2’ = ∆I’2’ = IO-IC1, with IC1 denoting the ionic current associated to the sub-state ‘C1’ and IO the ionic current through an open nanopore – substate ‘O’), and smaller than those of the PNA3-induced blockade events (Figure 4, panel c;) (∆IPNA3/IO = 0.995 ± 0.001, vide supra). To explain this, we remind that the bulkier PNA3DNA1 duplex (diameter of ~ 2.3 nm49) arrives at the β-barrel opening with the polypeptide tail head on, and is restricted

Figure 4. Free PNA3 and hybridized PNA3-DNA1 duplexes give distinct blockade signatures on the α-HL. (a) Selected recording reflecting the interaction of trans-added PNA3 (3 µM) with α-HL at ∆V = + 160 mV, in the presence of DNA1 (30 µM). (b) All-events histogram showing the distinct conductive states of α-HL corresponding to the putative nature of blockades shown in (a), as follows: free nanopore (O), the nanopore partly blocked by the PNA3-DNA1 complex (C1), and blockade associated to the passage of a free PNA3 molecule through the channel (U). (c) Representative view of a blockade event induced by unhybridized PNA3 interaction with the α-HL, still detectable in the presence of excess trans-added DNA1 (molar ratio 1:10). These events are distinct from those reflecting PNA3-DNA1 – α-HL interactions (events denoted by ‘1’ and ‘2’, panel d), whose amplitude correspond to the ‘C1’ sub-state in panel b. (d) Repre-



Page 6 of 10

(type ‘3’ events, panel b). Latter events, which duration are more frequent at higher applied voltages and are marked collectively as empty squares (□) in panel b, are indicative of the PNA3-DNA1 unzip. (c) Voltage dependence of the PNA3DNA1 unzipping events frequency (, ), corresponding to instances displayed collectively as ‘□’ in panel b, and the frequency of PNA3-DNA1 blockades not ending with the unzip ( ), denoted by ‘■’ in panel a. PNA3 and DNA1 were added to the trans-chamber in a molar ratio of 1:10. (d) The forces involved in the duplex unzipping, namely ( ), which acts on the cationic polypeptide attached to the PNA3 moiety, and a force acting oppositely on the negatively charged DNA1 strand ( ). (e) Beyond a threshold, the contribution of ( ) and ( ) leads to the unzipping of the PNA3-DNA1 duplex.

from fully entering the constricted space of the β-barrel. Assuming that a single amino acid residue from the polypeptide tail spans an axial distance of ~ 0.4 nm inside the nanopore, the 9-mer polypeptide tail attached to the PNA3 would accommodate entirely along the ~ 5 nm in length nanopore’s βbarrel, and give rise to the events denoted by ‘1’ and ‘2’ in Fig. 4, panel d. In contrast, the PNA3 monomer alone is capable of fully traversing the nanopore, thus giving rise to a slightly larger extent of block as the nucleic acid bases thread across the nanopore’s constriction region (Figure 4, panel c, or Figure 4, panel d, events denoted as type ‘3’ events). Unzipping a single PNA3-DNA1 duplex inside the α-HL The current levels denoted by ‘2’ and ‘3’ within the events depicted as ‘□’ in Fig. 5, panel b, were ascribed to the unzipping followed by translocation of a nanopore-captured PNA3DNA1 duplex, while the blockade events marked with ‘■’ reflect instances when the nanopore-captured PNA3-DNA1 duplex returns to the trans side without unzipping (Fig. 5, panel a). The PNA3-DNA1 duplex is prevented from entering the α-HL’s β-barrel, and the base-paired domain from the molecule can only interact at the mouth surface of the β-barrel (Fig. 5, panel d). Thus, the forces acting on the duplex have contributions from: (i) the electric force exerted on the PNA3 attached polypeptide tail ( ), which pulls it through the constriction region of the nanopore in the trans-to-cis directions and squeezes the duplex backbone against the β-barrel opening, and (ii) the electric force acting oppositely on the duplex from its negatively charged backbone from the DNA1 monomer ( ). As sketched in Fig. 5, panel e, once a threshold is reached, the unzipping is initiated, with the PNA3 monomer translocating the nanopore, and the DNA1 fragment returning to the trans side of the membrane, as dictated by the sign of the applied transmembrane potential.

This scenario is well supported by data shown in Fig. 5, panel c, which demonstrate that the frequency of the successful (‘□’) and respectively unsuccessful (‘■’) unzipping events increases, and respectively decreases, with the increase in ∆V. Note that the average time (̅ ) measured from durations associated to either successful unzip transitions of a captured PNA3 DNA1 duplex (type ‘2’ events, with life-times denoted by within ‘□’ events, Fig. 5, panels b) or unsuccessful unzips of a captured duplex (type ‘1’ events with life-times denoted by , marked with ‘■’ in Fig. 5, panels a), showed a nonmonotonic dependence with the applied potential (Fig. 6, panel a).

Figure 6. Statistical analysis of the dwell-times characterizing the molecular events preceding and subsequent to the PNA3DNA1 unzip. (a) Voltage dependence of the average dwell time ̅ measured on a population of time intervals comprising type ‘2’ events (Fig. 5, panel b) (successful unzip events) or unsuccessful unzips of a captured duplex (type ‘1’ events marked with ‘■’ in Fig. 5, panel a). Data points were fitted according to the model described in Fig. 7 (see text). (b) Voltage dependence of the average time of transitions measured within ‘□’ events and denoted as ‘3’ in Fig. 5, panel b, with durations, reflecting the PNA3 monomer threading across the nanopore following the PNA3-DNA1 duplex unzipping.

Figure 5. The voltage-induced unzipping of a single PNA3DNA1 duplex inside the α-HL. Shown are the blockade events associated to the PNA3-DNA1 capture inside the nanopore, seen as reversible, lower amplitude events of duration , which are more common at lower membrane voltages (∆V = +120 mV), marked as filled squares (■) (type ‘1’ events, panel a), and those displaying an initially lower ampli tude blockade step of duration (type ‘2’ events, panel b) assigned to a PNA3-DNA1 duplex captured inside the α-HL, followed irreversibly by a slightly larger blockade step of

The measured translocation time of the PNA3 strand following unzipping (transitions measured within ‘□’ events and denoted as ‘3’, with life-times in Fig. 5, panel b) shortened with the increase in the transmembrane potential (Fig. 6, panel b), as expected for a cationic analyte transported electrophoretically through the positively biased nanopore. Note however that such events measured at distinct ∆V’s, were significantly shorter as compared to the trans-to-cis transport


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


of PNA3 strands, seen in control experiments (Fig. S1, Supporting Information). The simplest explanation is that upon unzipping, the single stranded PNA3 escapes the nanopore by traveling a shorter distance, as measured from the constriction region towards the α-HL’s vestibule on the cis side (Fig. 5, panel e), as compared to the situation when the PNA3 translocations starts on the trans side (Fig. S1, Supporting Information).

transitions between discrete states and for the sake of completeness, we were able to rule out a more complex kinetic mechanism that included at least one more intermediate preceding the unzipped state. The strategy followed was to estimate the probability density function of blockade times intervals measured prior to the unzip, i.e. type ‘1’ transitions with , generating ‘■’ events, characterislife-times denoted by tic of duplex dissociation from the pore without unzipping (Fig. 5, panel a) and type ‘2’ transitions with life-times denot ed by within ‘□’ events (events preceding unzipping, Fig. 5, panel b), for a model containing an additional closed substate between the ‘C1’ and ‘U’ (Fig. S2, Supporting Information). As we present in details, such a probability density function (Figs. S3 and S4, Supporting Information) differs from an exponentially decaying function, which is predicted by the oversimplified model presented in Fig. 7 (see also expression 6, Supporting Information). On the other hand, the experimentally observed distribution of PNA3-DNA1 duplexinduced blockade time intervals on the nanopore (the events denoted by and in Fig. 5, panels a and b) reflects exponential decaying functions (Fig. S5, Supporting Information). This observation prompted us to neglect more complicated kinetic models, as the one depicted in Fig. S2, to describe the mechanism of PNA3-DNA1 unzipping inside the α-HL. By using the probability distribution function of block ade times for events denoted by and within the framework of the model shown in Fig. 7 (expression 6, Supporting Information), it can be demonstrated the simple relationship between the average value of such blockade intervals, and the corresponding rate constants for dissociation, of a system starting in the ‘C1’ sub-state: ̅ .

The kinetic and thermodynamic model describing the PNA3-DNA1 duplex unzipping inside the α-HL To explore the kinetic model describing the duplex unzipping inside the nanopore, we started by hypothesizing a two-step mechanism (Fig. 7).

Figure 7. Kinetic model describing the PNA3-DNA1-α-HL interactions at positive potentials. The nanopore found initially in an open state (O) (panel a), undergoes a transition to the closed state (C1) upon capturing the poly-arginine tail of the PNA3 from the duplex inside the lumen (panel b), with the association rate constant . From this sub-state (C1), the captured PNA3-DNA1 may dissociate and return to the trans side, with the dissociation rate constant , or unzip, with the DNA1 fragment returning to the trans side and the PNA3 monomer fully translocating the membrane (sub-state U, panel c). The unzip event is considered irreversible and characterized by the reaction constant rate . Depending on the value of the applied potential, the poly-arginine tail attached to the PNA3 moiety may not (panel d) or may (panel e) protrude deeply inside the lumen, under the influence. This results in a voltage-dependent value of the electric charge from the poly-arginine tail which gets harbored inside the lumen (see also text).

According the Kramers barrier theory, the PNA3-DNA1 duplex unzip rate constant ( ) vs. ∆V varies as ∆ 0!"# $

∗ ∆%&',

) *

+, where 0

) * ,

!"# $-

∗ ∆%.,

) *

+ gives

the thermal value of the unzip rate constant, measured in the ∗ absence of the applied potential, ∆01, is the standard free energy barrier for unzip in the absence of the applied potential, kB and T represent the Boltzmann constant and absolute tem∆

∗ perature, respectively. The term ∆0, = 2 3 ∗ , repre sents the supplementary quantity brought to the standard free energy barrier for unzip, associated to the movement of the duplex along the reaction coordinate during the unzip process inside the voltage-biased nanopore, qeff represents the effective electric charge from the duplex which interacts with the ∆V to determine the unzipping force, l is the nanopore’s length and 3 ∗ is the distance to the peak of the free energy barrier by which the duplex is separated along the reaction coordinate for unzip to occur.50 As presented in Fig. S6, Supporting Information, and with reference to the unzip step, larger ∆V values result in larger electric forces acting on the duplex, which determine lower values for the overall free energy barrier ∗ ∗ ∗ (∆0 ∆01, - ∆0, ) to be overcame by the duplex before the unzip occurs, and correspondingly larger unzip rate constants.

The duplex reversible association to the nanopore and irreversible unzipping inside the nanopore, can be empirically described as a barrier activation process within the Kramers rate theory in constant-force experiments.32-34 (see Supporting Information). It could be argued that the proposed scheme in Fig. 7 is oversimplified, since the molecular transition of the duplex to its unzipped state may contain intermediate states corresponding to conformations whereby the molecules finds the optimal conformation required to unzip, or the unzipping process itself may be described as a multi-step, consecutive reaction reflecting the step-by-step dehybridization of individual base-pairs. By employing the general Markov theory of

As for the step characterizing the duplex dissociation without unzip, it should be noted the net negative charge on the DNA1



∗ ∗ as compared to ∆01, , reflects the fact larger value for ∆01, that the unzip process requires more energy to occur as compared to the duplex dissociation without unzip.

favors it, while the net positive charge on the PNA3 hinders the duplex movement. Taking into account that the electric field is far larger inside the nanopore’s conductive pathway – where the PNA3 from the captured duplex resides - as compared to regions outside the lumen, where the DNA1 from the capture duplex experiences the electric field, it is safe to assume that with both electric contributions considered (i.e., from the PNA3 and DNA1 monomer), the dissociation process is energetically unfavored by ∆V. In other words, larger ∆V values result in larger values for the overall free energy barrier ∗ (∆0 ) to get overcome by the duplex before the escape to the trans side occurs, and correspondingly lower dissociation rate constants (k1O) (Fig. S6, Supporting Information).

CONCLUSIONS Herein, we demonstrated the uni-molecular discrimination between the un-hybridized and hybridized complexes of a cationic polypeptide-functionalized PNA with a ssDNA strand, by identifying and analyzing the distinct kinetics and amplitude of the ion current fluctuations ensued by their interaction with the α-HL. We then revealed the dissociation of PNA-DNA duplexes inside the voltage-biased nanopore, through the oppositely oriented electric forces acting on a cationic polypeptide tail attached to the PNA moiety, and the ssDNA backbone, respectively. From single-molecule experiments, we identified distinct sub-states and molecular pathways for PNA-DNA duplex capture, its escape without unzip, duplex unzipping and translocation, and measured the unzipping reaction rate constants and the free energy barrier height for the unzip. We demonstrated that the α-HL nanopore presents unique opportunities for detecting the hybridization at one end (vestibule) and capturing and unzipping the PNA3DNA1 duplex at the other (lumen). Based on this discovery, nanopore-based, force spectroscopy platforms may be further developed, enabling nucleic acids duplex detection and hybridization energy assessment during the same experiment. The presented approach may be useful when seeking: (i) diagnostic and accurate genotyping at the single-molecule level (ii) single-molecule methods to investigate metal-mediated base pairing in mismatched PNA-PNA or PNA-DNA complexes, to generate novel patterns of supramolecular assemblies, with controllable geometry and stoichiometry (iii) nanotechnological applications of PNAs for tailored designed force-sensing devices or real-time, molecular detection and recognition of biomolecules.

To describe the non-monotonic voltage-dependence of the ̅ values shown in Fig. 6, panel a, we assumed that the electric charge (qeff) from the PNA3-DNA1, sensing the electric field while plugging the nanopore, depends linearly with the ∆ (qeff ≈ 56. ∆). This is in line with previous findings,51 and reflects a scenario in which the larger is the applied potential, the larger becomes the extent to which the duplex crams inside the nanopore’s β-barrel (see also Fig. 7, panels d and e). With these considerations in mind, the unzip rate constant (k1U) varies vs. ∆V as ∆ 561!"#9∆ , while duplex dissociation rate constant (k1O) vs. ∆V varies as ∆ 562!"#-;∆ . The free parameters a and b contain lumped contributions from all terms entering the values of rate constants, as they are described within the Kramers barrier model being used (see above). The constants ct1 and ct2 are equivalent to the values of the rate constants associated to the unzip and duplex dissociation, respectively, measured in the absence of the ∆V (vide supra). As we show in Fig. 6, panel a, the non-linear fit with the equation 561!"#9∆ + 562!"#-;∆ , resulted in a good quantitative description of the average blockade time (̅ ) at different voltages, consistent with our experimental data. In the analytical expression of the rate constant for unzip at ∆V=0 (ct1), the function used for the fit analysis writes 561

) * ,

!"# $-

∗ ∆%.,

) *

ASSOCIATED CONTENT

+ (s-1). As stated previously for the

Supporting Information The Supporting Information is available free of charge on the ACS Publications website and contains additional experimental data regarding the un-hybridized PNA3 - α-HL interactions, as well as the detailed Markov model used to describe the kinetics of PNA3-DNA1 duplex - α-HL nanopore interactions.

Kramers model employed when diffusional transitions span ~ * nm distances, the Eyring frequency factor ( ) , where h is the , Planck constant, kB and T represent the Boltzmann constant and absolute temperature) equals 109 s-1.17 Combined with the values obtained from the fit (ct1=0.94), it follows that at a room temperature of T=300 K, the standard free energy barrier for the duplex unzipping in the absence of the applied poten∗ tial is ∆01, 11.6 59> ?@>! . Taking into account that in our experiment the PNA3-DNA1 duplex consists of 10 basepairs, and neglecting the small energy differences in pairing distinct bases, the rough standard molar energy for unzipping an individual base-pair equals 1.17 59> ?@>! , which is close to previously reported ones (1.47 59> ?@>! ).52 Similarly, knowing that on the model employed 562 ) * ,

!"# $-

∗ ∆%.,

) *

Page 8 of 10

AUTHOR INFORMATION Corresponding Author * [email protected] and [email protected]

Author Contributions #

These authors contributed equally

ACKNOWLEDGMENT

+ (vide supra), and considering that from the

non-linear fit in Fig. 6, panel a, ct2 = 2390.4, the value of the molar standard free energy barrier for duplex dissociation without unzip, in the absence of the applied potential, equals ∗ ∆01, 7.24 59> ?@>! . From the numbers above, a

8

We acknowledge the financial support offered by a National Research Foundation of Korea (NRF) grant funded by the Korean Government (No. 2016R1A2A1A05005440), Global Research Laboratory (GRL) Grant (No. NRF-2014K1A1A2064460) and Institute for Information & communications Technology Promotion (IITP) grant funded by the Korea government (MSIT) (No. 2017-0-01714), and Grant no. PN-III-P4-ID-PCE-2016-0026

ACS Paragon Plus Environment

Page 9 of 10 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 26. Schwarz, F. P.; Robinson, S.; Butler, J. M. Nucleic Acids Res. 1999, 27, 4792–4800. 27. Balagurusamy, V. S. K.; Weinger, P.; Ling, X. S. Nanotechnology 2010, 21, 335102. 28. Zhang, X.; Wang, Y.; Fricke, B. L.; Gu, L.-Q. ACS Nano 2014, 8, 3444–3450. 29. Wang, Y.; Tian, K.; Hunter, L. L.; Ritzo, B.; Gu, L.-Q. Nanoscale 2014, 6, 11372–11379. 30. Wanunu, M.; Morrison, W.; Kuhn, H.; Frank-Kamenetskii, M.; Meller, A. Nano Lett. 2010, 10, 738–742. 31. Tian, K.; He, Z.; Wang, Y.; Chen, S.-J.; Gu, L.-Q. ACS Nano 2013, 7, 3962–3969. 32. Mathé, J.; Visram, H.; Viasnoff, V.; Rabin, Y.; Meller, A. Biophys. J. 2004, 87, 3205–3212. 33. Sauer-Budge, A. F.; Nyamwanda, J. A.; Lubensky, D. K.; Branton, D. Phys. Rev. Lett. 2003, 90, 238101. 34. Dudko, O. K.; Mathé, J.; Szabo, A.; Meller, A.; Hummer, G. Biophys. J. 2007, 92, 4188–4195. 35. Hornblower, B.; Coombs, A.; Whitaker, R. D.; Kolomeisky, A.; Picone, S. J.; Meller, A.; Akeson, M. Nat. Methods 2007, 4, 315–317. 36. Tian, K.; Decker, K.; Aksimentiev, A.; Gu, L.-Q. ACS Nano. 2017, 28, 1204 – 1213. 37. Wang, Y.; Luan, B.-Q.; Yang, Z.; Zhang, X.; Ritzo, B.; Gates, K.; Gu, L.-Q. Sci. Rep. 2014, 4, 5883. 38. Schibel, A. E. P.; Fleming, A. M.; Jin, Q.; An, N.; Liu, J.; Blakemore, C. P.; White, H. S.; Burrows, C. J. J. Am. Chem. Soc. 2011, 133, 14778–14784. 39. Jin, Q.; Fleming, A. M.; Burrows, C. J.; White, H. S. J. Am. Chem. Soc. 2012, 134, 11006–11011. 40. Jin, Q.; Fleming, A. M.; Ding, Y.; Burrows, C. J.; White, H. S. Biochemistry 2013, 52, 7870–7877. 41. Johnson, R. P.; Fleming, A. M.; Jin, Q.; Burrows, C. J.; White, H. S. Biophys. J. 2014, 107, 924–931. 42. Perera, R. T.; Fleming, A. M.; Peterson, A. M.; Heemstra, J. M.; Burrows, C. J.; White, H. S. Biophys. J. 2016, 110, 306–314. 43. Ding, Y.; Fleming, A. M.; White, H. S.; Burrows, C. J. J. Phys. Chem. B 2014, 118, 12873–12882. 44. Asandei, A.; Chinappi, M.; Lee, J.-K.; Ho Seo, C.; Mereuta, L.; Park, Y.; Luchian, T. Sci. Rep. 2015, 5, 10419. 45. Chinappi, M.; Luchian, T.; Cecconi, F. Phys. Rev. E: Stat., Nonlinear, Soft Matter Phys. 2015, 92, 032714. 46. Montal, M.; Mueller, P. Proc. Natl. Acad. Sci. U. S. A. 1972, 69, 3561–3566. 47. Grosberg, A. Y.; Rabin, Y. J. Chem. Phys. 2010, 133, 165102. 48. Asandei, A.; Chinappi, M.; Kang, H.-K.; Seo, C. H.; Mereuta, L.; Park, Y.; Luchian, T. ACS Appl. Mater. Interfaces 2015, 7, 16706– 16714. 49. Eriksson, M.; Nielsen, P. E. Nat. Struct. Biol. 1996, 3, 410. 50. Nakane, J.; Wiggin, M.; Marziali, A. A Biophys. J. 2004, 87, 615– 621. 51. Heng, J. B.; Aksimentiev, A.; Ho, C.; Marks, P.; Grinkova, Y. V.; Sligar, S.; Schulten, K.; Timp, G. Biophys. J. 2006, 90, 1098–1106. 52. Ratilainen, T.; Holmén, A.; Tuite, E.; Nielsen, P. E.; Nordén, B. Biochemistry, 2000, 39, 7781–7791.

(NANOTWEEZ). This work was also supported in part by a grant of Ministry of Research and Innovation, CNCS - UEFISCDI, project number PN-III-P1-1.1-TE-2016-0508, within PNCDI III (Alina Asandei) and project number PN-III-P1-1.1-PD-20160737, within PNCDI III (Irina Schiopu).

REFERENCES 1. Bayley, H.; Cremer, P. S. Nature 2001, 413, 226–230. 2. Kasianowicz, J. J.; Balijepalli, A. K.; Ettedgui, J.; Forstater, J. H.; Wang, H.; Zhang, H.; Robertson, J. W. F. Biochim. Biophys. Acta, Biomembr. 2016, 1858, 593–606. 3. Gu, L.-Q.; Shim, J. W. Analyst 2010, 135, 441–451. 4. Howorka, S.; Siwy, Z. Chem. Soc. Rev. 2009, 38, 2360–2384. 5. Cao, C.; Long, Y.-T. Acc. Chem. Res. 2018, 51, 331–341. 6. Dekker, C. Nat. Nanotechnol. 2007, 2, 209–215. 7. Kowalczyk, S. W.; Blosser, T. R.; Dekker, C. Trends Biotechnol. 2011, 29, 607–614. 8. Kasianowicz, J. J.; Robertson, J. W. F.; Chan, E. R.; Reiner, J. E.; Stanford, V. M. Annu. Rev. Anal. Chem. 2008, 1, 737–766. 9. Single-Channel Recording, 2nd ed; Sakmann, B., Neher, E., Eds.; Springer US, 1995. 10. Asandei, A.; Mereuta, L.; Luchian, T. Biophys. Chem. 2008, 135, 32–40. 11. Song, L.; Hobaugh, M. R.; Shustak, C.; Cheley, S.; Bayley, H.; Gouaux, J. E. Science 1996, 274, 1859–1866. 12. Oukhaled, A.; Bacri, L.; Pastoriza-Gallego, M.; Betton, J.-M.; Pelta, J. ACS Chem. Biol. 2012, 7, 1935–1949. 13. Asandei, A.; Schiopu, I.; Iftemi, S.; Mereuta, L.; Luchian, T. Langmuir 2013, 29, 15634–15642. 14. Mereuta, L.; Asandei, A.; Seo, C. H.; Park, Y.; Luchian, T.. ACS Appl. Mater. Interfaces 2014, 6, 13242–13256. 15. Mereuta, L.; Schiopu, I.; Asandei, A.; Park, Y.; Hahm, K.-S.; Luchian, T. Langmuir 2012, 28, 17079–17091. 16. Wang, Y.; Gu, L. AIMS Mater. Sci. 2015, 2, 448–472. 17. Movileanu, L.; Schmittschmitt, J. P.; Scholtz, J. M.; Bayley, H. Biophys. J. 2005, 89, 1030–1045. 18. Reiner, J. E.; Balijepalli, A.; Robertson, J. W. F.; Campbell, J.; Suehle, J.; Kasianowicz, J. J. Chem. Rev. 2012, 112, 6431–6451. 19. Rosen, C. B.; Rodriguez-Larrea, D.; Bayley, H. Nat. Biotechnol. 2014, 32, 179–181. 20. Mereuta, L.; Roy, M.; Asandei, A.; Lee, J. K.; Park, Y.; Andricioaei, I.; Luchian, T. Sci. Rep. 2014, 4, 3885. 21. Asandei, A.; Schiopu, I.; Chinappi, M.; Seo, C. H.; Park, Y.; Luchian, T. ACS Appl. Mater. Interfaces 2016, 8, 13166–13179. 22. Angevine, C. E.; Chavis, A. E.; Kothalawala, N.; Dass, A.; Reiner, J. E. Anal. Chem. 2014, 86, 11077–11085. 23. Egholm, M.; Buchardt, O.; Christensen, L.; Behrens, C.; Freier, S. M.; Driver, D. A.; Berg, R. H.; Kim, S. K.; Norden, B.; Nielsen, P. E. Nature 1993, 365, 566–568. 24. Marchelli, R.; Corradini, R.; Manicardi, A.; Sforza, S.; Tedeschi, T.; Fabbri, E.; Borgatti, M.; Bianchi, N.; Gambari, R. In Targets in Gene Therapy; You, Y., Ed.; InTech: Rijeka, 2011; Chapter 2, pp 2947. 25. Tomac, S.; Sarkar, M.; Ratilainen, T.; Wittung, P.; Nielsen, P. E.; Nordén, B.; Gräslund, A. J. Am. Chem. Soc. 1996, 118, 5544–5552.

Authors are required to submit a graphic entry for the Table of Contents (TOC) that, in conjunction with the manuscript title, should give the reader a representative idea of one of the following: A key structure, reaction, equation, concept, or theorem, etc., that is discussed in the manuscript. Consult the journal’s Instructions for Authors for TOC graphic specifications.

Insert Table of Contents artwork here



TOC 39x23mm (300 x 300 DPI)

ACS Paragon Plus Environment

Page 10 of 10

Single Molecule, Real-Time Dissecting of Peptide Nucleic Acids-DNA

Recommend Documents