Conformational Changes Followed by Complete Unzipping of DNA

Subscriber access provided by GAZI UNIV

Article

Conformational Changes Followed by Complete Unzipping of DNA Double Helix by Charge-Tuned Gold Nanoparticles Subhas Chandra Bera, Kasturi Sanyal, Dulal Senapati, and Padmaja P Mishra J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.6b01323 • Publication Date (Web): 15 Apr 2016 Downloaded from http://pubs.acs.org on April 16, 2016

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.

The Journal of Physical Chemistry B 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 28

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 Journal of Physical Chemistry

Conformational Changes Followed by Complete Unzipping of DNA Double Helix by Charge-Tuned Gold Nanoparticles Subhas C. Bera,†,# Kasturi Sanyal,†,# Dulal Senapati† and Padmaja P. Mishra*,† †

Chemical Sciences Division, Saha Institute of Nuclear Physics, Kolkata, India

*Corresponding author Email:[email protected]

1 Environment ACS Paragon Plus


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: The complete unzipping of DNA double helix by small size gold nanoparticles having weakly positive surface charge has been monitored using ensemble and single molecule fluorescence resonance energy transfer (smFRET) techniques. We believe, as the gold nanoparticles have positive charge on its surface, the DNA and nanoparticles were pulled together to form two single strands. The positively charged ligands on the nanoparticles attached to the DNA, and the hydrophobic ligands of the nanoparticles became tangled with each other, pulling the nanoparticles into clusters. At the same time, the nanoparticles pulled the DNA apart. The conformational changes followed by unzipping have been investigated for both long DNA (calf thymus DNA) as well as for short DNA (~40 base pair) using ensemble methods like circular dichroism (CD) spectroscopy, fluorescence intercalation assay, viscometric and single molecule FRET imaging. This observation not only reveals a new aspect in the field of nano-bio interface but also provides additional information about DNA dynamics. 1. INTRODUCTION The nature and strength of interaction between DNA and its components with synthetic molecules, capable of DNA-binding have resulted in the development of several classes of designed scaffolds accounting to their unique physico-chemical properties.1,2 Alternate approaches in the search for molecules appropriate for interaction with DNA sequences have procreated in the identification of systems capable of covalent modification of the target DNA strand.3,4 DNA and its synthetically programmable sequence recognition properties have been utilized to assemble nanoparticles functionalized with oligo nucleotides into preconceived architectures.5-8 The widely studied gold nanoparticles (AuNPs) represent an admirable biocompatibility9,10 and hence, the concoction of AuNPs with biomolecules find a wide range of


Page 2 of 28

Page 3 of 28

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


application from nanoelectronic devices11,12 to biomedical applications including diagnostic and therapeutic treatment.13-15 The DNA conformation (mainly the nature of helicity i.e. A, B or C form DNA), chain rigidity (i.e. the nature of nucleotide sequences), size and charge of nano particles plays an imperative role in their ability to induce the interaction with DNA, through a combination of electrostatic attraction,16 groove binding,17 and intercalation. Such binding events induce the changes in the conformation of the DNA strand, and for each AuNPs-DNA system, a global effect is exerted in conjugation with both the metal cluster and the capping agent of the nanocluster. If the nanoparticles have highly positive surface charge density, the DNA is likely to wrap and bend upon binding to the nanoparticles.18 On the other hand, it has been observed that the rate of adsorption of shorter DNA strands on the AuNPs is faster compared to that of longer strands, where as the longer DNAs bind more tightly to the AuNPs due to the establishment of more contacting points on the AuNP Surface.19 In a similar note, due to a combination of electrostatic repulsion and dipolar interaction, ssDNA preferably adsorbs on the surface of AuNPs compared to dsDNA20,21, which is further facilitated in presence of salt.22 Nonspecific binding of ssDNA to gold nanoparticles is strongly dependent on the ionic strength of the medium and can be suppressed by using sufficiently high concentration of electrolyte. However, for dsDNA, the hydrogen bonding between the two strands plays a crucial role to determine the extent of interaction. A number of studies have also reported the chemisorptions of oligos on the AuNP surface resulting conformational change in the DNA.19,23 So far, majority of the research in this field have been focused on negatively charged AuNPs.24 However, cationically modified AuNPs become an attractive and rather obvious choice for DNA interaction, particularly in view of the several advantages.25,26 Moreover, understanding of the



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

non-covalent interaction of the semi flexible DNA with AuNPs is required to develop the nanoparticles applications. It still remains unclear if the DNA is stable enough in aqueous solution when interact with AuNPs, which could be a crucial point for applications of the action of oligo nucleotide-AuNP conjugates. In this work, we have used small sized (4 nm-10 nm) positively charged AuNPs bearing nearly one tenth of the surface charge density of a histone octamer, to study the interaction with dsDNA. The motivation is to define the relative importance of nanoparticle and experimental variables that contribute to the observed conformational changes and stability of DNA. Our studies show a captivating observation that tuning the size and charge of the AuNPs results in a conformation change followed by a completely unzipping of the dsDNA. This is in contrary to the results in this field, as most of the works reported the adsorption of DNA on gold surface rather than unzipping.23 Recently, Melechko et al. has reported that weakly charged gold nanoparticles induce strand separation in dsDNA, giving an preliminary observation that provide insight into the interaction of dsDNA with those tuned AuNPs.17 Here we have suggested a detailed mechanism using ensemble and single molecule fluorescence measurements to ensure the strand separation. 2. EXPERIMENTALS 2.1. Materials: All chemicals used are of analytical grade if not otherwise stated are purchased from SigmaAldrich and used with no further purification. The HPLC purified Cy3 and Cy5 labeled oligo were purchased from IDT, USA and used as received without any further purification. Salts used to make buffer for the experiment were purchased from Merck. All solutions were prepared using mili-Q water and sterilized by autoclave or 0.22 µm membrane filter.


Page 4 of 28

Page 5 of 28

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


2.2. Preparation of surface modified AuNP: A slightly modified protocol described by Grueso et. al. has been used to synthesize the nanoparticles.27 In brief: 2 ml of 10 mM HAuHCl4.3H2O was mixed slowly with 11 ml of 10 mM aq. L-cysteine solution with constant stirring at room temperature. 240 µL of 100 mM aq. NaBH4 (freshly prepared with cold water) was then added drop wise to the mixture. After the complete addition, reaction mixture turned ruby red. The mixture was incubated in ice for 30 minutes with stirring. The concentration of the AuNPs was calculated from the estimated extinction coefficient and by using the subsequent Beer-Lambert’s Law. From the reported results28 of extinction coefficient of different spherical particles with different absorption maximum, we have constructed a linear plot. Extrapolation of this linear plot to the desired wavelength of the L-cysteine capped gold nanoparticle gives us the measure for the extinction coefficient from where we back calculated the concentration of the used nanoparticles. 2.3. Transmission electron microscopy (TEM): For TEM micrographs, a single drop of the aqueous solution (0.1 mg ml-1) of the gold nano particles was placed on a carbon coated copper grid. The grid was left to dry for several hours at room temperature inside desiccators. TEM images are taken using a Philips CM 200 electron microscope working at 200 KeV. Size distributions of the Au core were measured from enlarged TEM images for at least 200 individual cluster cores. Calculation of concentration of the AuNP was done following previous repots.29 2.4. Zeta potential experiment: Zeta potential measurement was carried out with a ZEN 3690 Zetasizer Nano ZS90 model. Zeta dip cell was used at 25 °C. The zeta potential distributions of AuNPs measured as 8 mV, 14 mV and 22 mV for 4 nm, 6 nm and 10 nm AuNPs respectively in water. The charges on the AuNPs



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

are sufficient to avoid inter particle interaction and thus maintaining a stable particle size for a week, at least. Hence all the experiments were carried out within 3-4 days of preparation of the AuNPs. 2.5. Circular dichroism spectroscopy: Electronic circuler dichroism (CD) spectra were recorded in a Chirascan spectropolarimeter (Applied Photophysics, UK). A standard quartz cell of 1 mm path length was used. The CD spectra were taken at fixed calf thymus DNA (ctDNA) concentration and varying the concentration of the AuNPs of different size in different experiments. The spectra were expressed in terms of molar ellipticity. Scans were taken from 220 nm to 320 nm for the intrinsic region. For each spectrum, three consecutive readings were averaged at a constant temperature of 298 K with a 5 min equilibration before each sample. 2.6. Ethidium bromide (EB) displacement assay: Before measurement, DNA (20 µg/ml) was incubated with EB (0.8 µg/ml) for 1 hour. The AuNPs of different size then titrated with the EB-DNA solution separately and the fluorescence was measured by exciting at 520 nm at temperature of 25°C using Fluoromax-3 (Jobin Yvon, Horiba scientific, Japan) spectrofluorimeter. 2.7. Viscosity measurement: Viscosity experiments were performed at 25°C immersing in a thermostatic water bath. Various amounts of AuNPs of different size were then added to a solution of DNA in the viscometer while keeping the DNA concentration constant. The flow times of the samples were repeatedly measured with an accuracy of ± 0.2 s by a digital stopwatch. Each point was the average of at least three measurements. The buffer flow time (t0) was observed, and relative viscosities for DNA in the presence and absence of AuNPs were calculated from the relation: η = (t – t0)/t0,


Page 6 of 28

Page 7 of 28

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


where t is the observed flow time. The data were presented as plots of (η/η0)1/3 versus [AuNPs], where η and η0 are the viscosity of DNA in the presence and absence of AuNPs, respectively. 2.8. Labeled DNA substrate: High-pressure

liquid

chromatography

(HPLC)

purified

oligo

nucleotides

(5′-Biotin-

TGGCCAAAAAAGCATTGCTTATCAATTTGTTGCACCGACCCTA-A-Cy3, and 5’-Cy5TTAGGGTCGGTGCAACAAATTGATAAGCAATGCTTTTTTGGCCA-3′) were purchased from Integrated DNA Technologies. To anneal for making the dsDNA, equimolar amounts of the above primers, made with 10 mM Tris-HCl, pH 8.0 and 50 mM Mg2+ buffer, were mixed and heated at 92°C for 4 min and gradually cooled down to room temperature. 2.9. Steady-State absorption and fluorescence spectroscopy: Absorption measurements were carried out in a JASCO V-650 UV-Vis Spectrometer. Two standard quartz cuvette of 10 mm path length were used. Scans were taken from 350 to 750 nm at 298 K. Fluorescence measurements were carried out at 298 K in a Hitachi F-7000 spectrofluorimeter. The experiments of comparative conjugation were done by adding different gold nanoparticle concentration. The excitation and emission wavelength were 545 and 555 nm. The emission was collected using slit width 5 nm and PMT voltage of 900V. 2.10. Time resolved fluorescence: Time-resolved emission spectra were recorded using a MCP-PMT detection based picoseconds pulsed diode laser integrated TCSPC ﬂuorescence spectrometer (Jobin Yvon, Horiba scientific, Japan) with λex 471 nm, λem 564nm and 661 nm for Cy3 and Cy5 respectively. The emission from the samples was collected at a right angle to the direction of the excitation beam. The full width at half maximum (FWHM) of the instrument response function was 250 ps and the resolution was 28 ps per channel. The data were fitted to multi-exponential functions after



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

deconvolution of the instrument response function by an iterative reconvolution technique using IBH DAS 6.2 data analysis software in which reduced χ2 and weighted residuals serve as parameters for goodness of fit. 2.11. Single-Molecule FRET experiment: We used a standard procedure30 to make the dsDNA from single stranded-oligos (ss-oligos) having Cy3 or Cy5 attached at one end. PEG/Biotin-PEG-coated predrilled quartz microscope slides were used to monitor fluorescence signals from single Cy3 and Cy5 fluorophores. Cy3attached DNA substrate was immobilized on a quartz slide through biotin–streptavidin interaction. The surface modified AuNPs were delivered to the premade channels containing the DNA substrate. A 532 nm laser was used to excite the Cy3, and then the fluorescence intensities from Cy3 and Cy5 were recorded simultaneously using a prism-type total internal reflection (PTIR) based inverted microscopic system. FRET time traces were analyzed with HMM using a home-built HMM optimization code. Our instrumental setup is shown in Scheme S1, has been constructed on an inverted microscope (Olympus IX 71). A Solid state 532 nm Diode laser (Laser quantum, UK) was used to excite Cy3 by total internal reflection to reduce the background signal from solution. The emission signal was collected by a water immersion objective (60x, 1.2 NA, Olympus) and then split into donor and acceptor channels using a 550-630 nm dichroic mirror (640DCXR, Chroma) after passing through a long-pass filter (550 nm, Chroma). FRET time traces were recorded using a electron multiplying charge coupled device (emCCD, Ixon3+ 897, Andor Technologies, South Windsor, CT) camera with image integration time of 30 ms. The data acquisition software used was the generous gift from Tae-Hee Lee (Pensylvania State University, USA) or T. J. Ha (University of Illinois, USA), developed using Visual C++ (Microsoft, WA). The raw data were pre-analyzed


Page 8 of 28

Page 9 of 28

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


by a program written in IDL, with which FRET spots were extracted from the raw data. Bleed though of Cy3 emission to the Cy5 channel is typically < 5% of the Cy3 emission, and is subtracted from the Cy5 emission during data analysis. Direct excitation of the Cy5 by the laser was almost negligible. All experiments were done at room temperature unless otherwise specified. The FRET efficiency was calculated after correction for the cross talk between the donor and acceptor channels. In order to suppress the noise, five adjacent data points in a single-molecule time trajectory were averaged before further FRET analysis unless otherwise specified. Individual traces were chosen where donor and acceptor exhibit visible clear anti-correlation and one-step photobleaching, which indicates one dye per DNA. FRET histograms were built by using the regions free of blinking and photobleaching and combining molecules at equal contribution per molecule. Cross-correlation curves between the donor and acceptor trajectories were calculated. To obtain the statistics of DNA’s diffusion dynamics, more than 100 molecules were included in the analysis of both FRET histograms and cross-correlation curves. The FRET efficiency was computed using the relationship: ‫ܧ‬௘௙௙ = (ூ

(ூ಴೤ఱ ) ಴೤ఱ ାூ಴೤య )

.............................................................................................................................1

Where Eeff is FRET efficiency and Icy3 and Icy5 are intensity of Cy3 and Cy5 respectively. Corresponding FRET distance was calculated using the following equation: ோల

బ ‫( = ܧ‬ோల ାோ ల ) ......................................................................................................................................2 బ

Where, R0 and R were the Förster distance and distance between the donor-acceptor respectively. HMM algorithm was used to extract all the kinetic and other analytical information from the raw smFRET traces. The details of the HMM can be found elsewhere.31



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

Page 10 of 28

3. RESULT AND DISCUSSION 3.1. Fluorescence Displacement Assay: The fluorescence displacement assay has been carried out to get a preliminary estimation of how the initial DNA conformation influences the course of AuNP binding.32 The competitive binding of the AuNPs with EB reflected a decrease in the fluorescence intensity of the EB, followed by complete quenching. The mode of interaction between AuNPs and DNA has been found to depend not only on the [AuNP]:[DNA] ratio, but also on the size and hence the charge of the AuNPs (as size difference leads to change the number of ligands on its surface resulting variation in surface charges). Fig 1(A) clearly indicates that the efficiency of displacement of intercalated EB is more for 4 nm AuNPs (surface charge + 9 per nanoparticle), compared to that of 10 nm AuNPs (surface charge + 22 per nanoparticle). The equilibrium binding constant found to be 8.4 × 106 M-1 for the AuNPs with 4 nm diameter and the same for the 10 nm AuNPs is calculated to be 3.6 × 106 M-1. Though, the result indicates displacement of EB that does not mean competitive intercalation only but can also indicate removal of EB due to DNA conformation change which is efficiently higher for smaller AuNPs. To clarify the actual effect further studies were performed. 3.2. UV/Vis centrifugation assay: The stoichiometry of the DNA-nanoparticle interaction was established via a UV/Vis centrifugation assay.33 In this assay, the addition of an agent that binds DNA and converts the coiled strands into condensed strands, which precipitate out with centrifugation. Determination of the critical mass of the condensing agent indicates the binding ratio of the two molecules. From this, it was established that the AuNP:DNA ratio is 7.8:1, 5.4:1 and 3.2:1 for AuNPs of 4 nm, 6 nm and 10 nm respectively. This AuNP:DNA ratio appears to be logical based on the


Page 11 of 28

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


relative length scales of the two molecules. For example, the DNA strand is approximately 14.3 nm in length, while each small sized AuNP is about 4 nm, suggesting four nanoparticles could bind to each “side” of the DNA strand. 3.3. Circular dichroism spectroscopy studies: The dynamic change to the DNA structure caused upon the interaction with the AuNPs during the course of the binding has also been followed by the circular dichroism spectroscopy (Fig.1B&C). As is known, the backbone conformation of DNA shows a CD spectrum characteristic of the right-handed B form in the far UV region (220–320 nm). Structural alterations of the DNA caused by its interaction with AuNPs are reflected in changes in this intrinsic CD spectrum. The CD spectrum of free DNA has a positive peak at approximately 278 nm (right arrow) and a negative peak at 247 nm (left arrow) which corresponds to B-DNA in Fig. 1(B). These are caused by stacking interactions between the bases and the helical suprastructure of the polynucleotide that provides an asymmetric environment for the bases.34 With increase in the concentration of the AuNPs, the ellipticity (θ) of the positive peak decreased, whereas, that of the negative peak increased for all the three different size AuNPs used for the experiment, followed by a change in the λmax position at the positive one (middle arrow). This indicates the removal of H-bonds as well as partial denaturation of the double helix. This is because, groove binding and electrostatic interactions are assumed to show no or little perturbation on either of the peaks, while intercalation enhances the intensities of both peaks.27 Strand elongation, on the other hand, has been assigned to increase in the molar ellipticity of positive and negative peaks.35 The shifting of the λmax position is an indication of formation of simple charge pairing of the cationic AuNPs with the phosphate back bone of the DNA. The effect of L-cysteine and pH was negative on calf thymus DNA (ctDNA). Fig. 1(C) shows the relative molar ellipticity at the



100

4 nm AuNP 6 nm AuNP 10nm AuNP

60 40 20 0

4 θ (mdeg)

Fl.Int.(%)

80

0 -4 -8

(A)

260 280 360 λ (nm) 4 nm AuNP 6 nm AuNP 10 nm AuNP

1.0 0.9 0.8

0.3 0.0

d e f g

(η/η0)

0.6

240

1/3

4 nm AuNP 6 nm AuNP 10 nm AuNP

0.9

a b c

(B)

1E-4 1E-3 0.01 0.1 [AuNP]/[DNA]

[θ]/[θ0] 280 nm

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

Page 12 of 28

0.7

(C)

0.00

0.6 0.04 0.08 [AuNP]/[DNA]

0.12

(D)

0.0

0.4

0.8 1.2 1.6 [AuNP] µM

2.0

Figure 1: (A) EB fluorescence displacement assay with different sized AuNPs. (B) CD spectra of ctDNA at different concentrations of 4 nm AuNP (0 µM (a), 2 µM (b), 4 µM (c), 5 µM (d), 10 µM (e), 20 µM (f) and 100 µM (g). (C) Effect of AuNP size on CD of ctDNA. (D) Change in viscosity of ctDNA in presence of different size AuNPs. maximum of the positive peak proportionate to the change in the AuNP size. However the extent of change in molar ellipticity at a particular concentration of AuNP has been observed to depend on the size of the corresponding AuNPs. We had an interesting observation showing both the peaks almost diminished with increase in the concentration of 4 nm AuNP. However, the peaks did not vanish completely for the other two AuNPs (Fig. 1B & C). The higher binding constants obtained from Chipman plot (Fig S3) also indicates greater affinity of DNA towards the AuNP. 3.4. Viscosity measurements:


Page 13 of 28

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 relative viscosity of the DNA has been observed to decrease with increase in AuNP concentration (Fig. 1D). The results correlated well with that of reported previously to investigate induced structural changes in the DNA double helix by AuNPs.36 The decrease in the viscosity for all the three different size AuNPs, indicating the binding mode that induces a DNA conformational change to a less elongated form, i.e. the DNA undergoes a structural deformation (sec 3.3) followed by formation of a compact structure around the AuNPs due to adsorption on the AuNP surface. A groove binding would result in no change in the viscosity of the solution, as there is no change in the length of the DNA helix.27 Like the above experiments, the 4 nm AuNPs induced a substantial decrease in the viscosity compared to the bigger one (10 nm) at a particular concentration. The measured zeta potential (data not shown) was also higher, once the relative viscosity of the system is lower, because of the shrinkage of the shear plane that would suppose a more compact DNA/AuNP conformation. The experiments so far were carried out with ctDNA, and the results clearly explain a loss of the double helix of the DNA, probably a breakage of the phosphate backbone and formation of compact structures around the AuNPs. Moreover, one significant factor to be deliberated about is, whether the DNA forms the compact structure as being in the duplex form (i.e. the rigid form) or undergoes a strand separation to form ssDNA before forming a compact structure with the AuNPs. It also remains unclear if these AuNPs are effective enough to induce similar changes in smaller DNAs. To find out the plausible hints we have carried out the experiments with synthetic oligos labeled with Cy3 and Cy5 as FRET donor–acceptor pair at two ends (5΄-BiotinTGGCCAAAAAAGCATTGCTTATCAATTTGTTGCACCGACCCTAA-Cy3, TCGGTGCAACAAATTGATAAGCAATGCTTTT-TTGGCCA) efficiency while hybridized.


having

and

maximum

5΄-Cy5FRET


3.5. Steady-State absorption and fluorescence studies: The effect of AuNPs on the absorbance has shown a decrease at 550 nm (Cy3 absorption maxima) as well as 650 nm (Cy5 absorption maxima) with increase in AuNP concentration (up

1600

(A)

Fl.Int. (A.U.)

90

Cy3

1200

(B)

0.9

E(fret)

0.6 60

800

0.3

Cy5

400 0

30 0

1600

2

4 6 8 [AuNP] (µM)

0 1500

2

4 6 8 [AuNP] (µM)

(D)

10 µM AuNP

10

Fl.Int. (A.U.) Linear fit

1250

800

1000 60 µM AuNP

400 0

0.0

10

(C)

1200

Fl.Int. (A.U.)

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

Page 14 of 28

560

600

640

680

750 10

20

λ (nm)

30 40 50 [AuNP] (µM)

60

Figure 2: Steady state fluorescence data. (a) Increasing Cy3 intensity along with reverse change of Cy5 one, indicates loose of FRET up to a certain AuNP conc. (b) Corresponding E(fret) plotted with respect to [AuNP]. (c) Quenching of Cy3 fluorescence intensity at higher [AuNP] and (d) the linear fit of it.

to 60 µM), irrespective of their sizes (Fig. S2). This could be because of the changes in the dipole orientation of the dyes upon adsorption on the surface of the AuNPs. This was also further verified by steady state FRET measurement. As the donor (D or Cy3) and acceptor (A or Cy5)


Page 15 of 28

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


were labeled at the same end of the dsDNA, the FRET efficiency was expected to be as high as 0.85, indicating an approximate distance of 40 Å between the FRET pairs. A substantial loss of FRET efficiency was observed with increase in the AuNPs concentration till 1.2 µM as reflected an antiphonal decrease in the Cy5 intensity with increase in Cy3 (Fig. 2A). The FRET efficiency was almost negligible for 4 nm size AuNPs, whereas for the 6 nm and 10 nm AuNPs some energy transfer efficiency was retained even at the higher concentrations (Fig. 2B). The decrease in FRET efficiency, irrespective of the size of AuNPs, argues an increase in the distance between DA pair caused by strand separation, as change in helicity or strand elongation are not expected to induce similar changes. Additionally with further increase in the AuNP concentration, the Cy3 intensity also experienced a decrease and finally quenched completely for all the AuNPs (Fig. 2C and D). This phenomenon could explain the adsorption of ssDNA onto the AuNPs surface resulting in static quenching of the fluorescence intensity.37 3.6. Time resolved fluorescence studies The time resolved fluorescence results also support the steady state fluorescence results (Fig. S4 and Table S1). The free Cy3 has a very short life time (τ) of approximately. 0.4 ns in agreement with its known low quantum yield (≤ 0.04). Cy3 when labeled with oligo displayed two lifetimes (amplitudes) of 0.32 ns (0.6) and 3.9 ns (0.4) as shown in Fig. S4. The double exponential life time distribution are likely due to a fast exchange between two conformational states of the dye coupled to DNA: a state where the dye is primarily surrounded by polar water molecule (0.32 ns) and a state in which the dye stays in close proximity to the less polar DNA (3.9 ns).38 Upon conjugation of the acceptor, Cy3 showed an additional life time. Cy3 is expected to have many relaxation pathways, considering the complexity in the life time. In addition, other complexions also appear as Cy5 is known to isomerize between a fluorescent trans and a non-fluorescent cis



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

Page 16 of 28

state,39 to have a significant triplet state in an oxygen depleted environment.40,41 The energy transfer between both triplet state and singlet state of donor to triplet state of acceptor are also possible, however it has been studied that these efficiencies are closely similar to the traditional singlet–singlet transfer processes.40,41 With increase in the concentration of the AuNPs, the longer life time value of Cy3 decreased from 3.9 nm to 2.2 nm for a concentration of 4 µM of AuNP followed by an increase of the value to 3.1 ns, whereas the shorter life time experienced a continuous 2.5 fold increase. However, both the τ values for Cy5 decreased with increase in the AuNP concentration, reflecting a decreasing the FRET efficiency, and hence an increase in the distance between the donor-acceptor pair. As a control we measured the τ values for the free donor, the free acceptor and the donor labeled ssDNA in both absence and presence of complementary DNA and dsDNA labeled with both donor and accepter at a close proximity. From the control experiment it was observed that the τ values at higher [AuNP] during titration closely resembled to that of the only donor labeled ssDNA in presence of AuNP. This also supports the formation of ssDNA after AuNP interaction compared to that of previous steady state fluorescence results. 3.7. Single-Molecule FRET studies: Although many existing methods characterize both biomolecules and nanomaterials, majority of them are unable to give a detailed picture of biomolecular structure at the nanomaterialbiomoclecule interface. This is becoming increasingly necessary to advance fields of research that hinge on nano-bio interactions. The existing methods often focus on a detailed understanding of either of the individual systems, but have insufficient overlap to study both simultaneously. Hence, the local electronic properties, bioavailability, toxicological effects, and basic molecular structure and conformation of biomolecules on nanoparticles remain unclear.


Page 17 of 28

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


This hampers the ability to predict relevant biological activity on nanoparticles, and leaves us illequipped to design sensors, biocatalysts, and medical diagnostic tools. Single molecule FRET is very appealing in this field because of its simpleness of building ratiometric fluorescent systems. Unlike those one-signal sensors, the ratiometric sensors contain two different chromophores and use the ratio of the two fluorescence intensities to quantitatively detect the analytes, and they can eliminate most ambiguities in the detection by self-calibration of two emission bands. Those external factors, such as excitation source fluctuation and sensor concentration will not affect the ratio between the two fluorescence intensities in single molecule measurements though a common characteristic of these approaches is that they limit the volume from which fluorescence signals are acquired.42,43 Due to various advantages of this single molecule FRET technique, this has been used an important tool to study such complex biological phenomena with great findings. The position of the donor–acceptor pair along the length of the oligo nucleotides have been examined to measure the FRET response, and it was found that the three dimensional context of inter nucleotide-tethered fluorophores have a direct effect on the strength of the FRET signal. Our attempt of using single molecule FRET imaging has provided information that would rather remain difficult to be observed from ensemble measurements. To perform the single molecule FRET study (Fig. 3), we used PEG/Biotin-PEG-coated predrilled quartz microscope slides to monitor fluorescence signals from single Cy3 and Cy5 fluorophores. Cy3-attached DNA substrates have been immobilized on a quartz slide through biotin– streptavidin interaction. The surface modified AuNPs were delivered into the premade channel containing the DNA substrate. A 532 nm LASER has been used to excite Cy3-labeled DNA substrate and fluorescence intensities from Cy3 and Cy5 were recorded simultaneously using a prism-type total internal reflection (PTIR) based inverted microscopic system (Scheme S1).



FRET time traces (Fig. 3(A) and (D)) were analyzed using a Hidden Markov model (Fig. 3(B) and (E)) by a HMM optimization code received as a generous gift.31

1.2

Real time delivery of AuNPs Cy5 Cy3

2500

1.2 E(fret) HMM fit 0.8

0.8 E(fret)

Fl.Int. (A.U.)

5000

# events

0.4

0.4

0.0 0.0

0 0

5

10

15

20

0

3

6

9

1.2

Real time delivery of AuNPs

2500

0.8

15

Time(Sec)

20

30

45

# events

0.0

0.0

0

(D)

15

0.4

0.4

10

0

(C)

E(fret)

Cy5 Cy3

5

15

1.2 E(fret) HMM fit 0.8

5000

0

12

(B)

(A)

Fl.Int. (A.U.)

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

Page 18 of 28

0

3

6 9 12 Time(sec)

0

(E)

15

30 45 # events

60

(F)

Figure 3: Single-Molecule FRET experiment with labeled oligo. (A) Real time FRET time traces, (B) HMM analysis & (C) population distribution histogram obtained after analyzing the effect of AuNP to end–labeled oligos. (D), (E) & (F) are the real time FRET time traces, HMM analysis & population distribution histogram respectively showing the effect of AuNP to mid– labeled oligos. Both of them are indicating two distinct intermediate FRET states.

We have observed a initial high FRET state of about 0.82 (Fig. 3(B) and (E)) from the surface immobilized DNA, as the donor-acceptor pair are labeled on the same site (5΄ of one oligo and 3΄ of the complementary oligo) with a six nucleotide overhang of the dsDNA, supporting a distance


Page 19 of 28

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 about 3-4 nm between the donor-acceptor pair. However, the FRET value decreased gradually in a step wise manner, after addition of the AuNPs to the immobilized DNA followed by complete decrease of the FRET abruptly (Fig. 3). In the meanwhile, a slight increase in the Cy3 intensity is also monitored in many cases, followed by a decrease as well. For repetition of the same experiment with different size AuNPs, nearly similar results have also been observed. Be that as it may, when the experiments were carried out with mid labeled dsDNA (donor and acceptor are labeled at the mid position of the individual oligo and then annealed) a delayed but similar response has also been encountered (Fig. 3(A) & (D)). The HMM analysis of the smFRET results clearly indicate the existence of two distinct intermediate FRET states with Efret values 0.67, 0.39 and a substantial contribution of the zero FRET state (Fig. 3(B) and (E)). The individual FRET states could be assigned to initial binding of the AuNPs with oligos, a transition state of higher energy followed by a complete strand separation. The average strand separation time is 23±3 s for the AuNPs of 4 nm where as it increased till 31±2 s for that of 10 nm size. In a control experiment of adding AuNPs to ss-oligos labeled with Cy3 only at the 5΄ end or mid position experienced a quenching of the fluorescence intensity with in a fraction of second. We did not encounter step wise quenching at single molecular level, indication the stepwise decrease of the FRET values in case of the dual labeled double stranded-oligos (ds-oligos) are assuredly due to the interaction induced of the AuNPs with them. The experimental results presented here, clearly indicate that the addition of AuNPs to the dsDNA changes the structure, thus affecting the helicity of the DNA followed by a separation of the strands. This could be due to several collective effects for both long as well as short DNAs. The synthesis procedure of the AuNPs and the Zeta potential measurements argues that the



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

Page 20 of 28

surface charge of the particles to be positive in biological condition. Thus during the interaction of these

Scheme 1: Representation of the plausible mechanistic way of dsDNA unzipping in presence of surface modified AuNP. The upper panel represents the complete unzipping of the dsDNA after experienced a conformational change due to interaction with the 4 nm AuNPs. The lower panel represents the conformational changes and partial unzipping of dsDNA, when interacts with the bigger size (10 nm) AuNPs.

AuNPs with the negatively charged phosphate group of the dsDNA, the synergistic role of charges and hydrophobicity induces the temporal loss of WC hydrogen bond network as well as


Page 21 of 28

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


base pare stacking. However, it also suggests that particles acting in concert could only induce the structural changes, which are not believably possible for single AuNPs. Comparing the shape and size of DNA to that of AuNPs used, it is most unlikely for the nanoparticle to directly insert to the base pairs through either end of the DNA, or into the grooves without inducing conformational alternation. This overall gives us a mechanism, which includes a number of phenomena, namely DNA compaction with addition strand separation, DNA bending, to name few (Scheme 1). However, more experiments need to be carried out to confirm and characterize the two intermediates to provide a detailed mechanism.

4. CONCLUSION This work provides insight into the interaction of dsDNA with surface modified gold nanoparticles bearing very less surface charge density compared to a histone octamer. We have observed a collaborative effect of the nanoparticles resulting structural changes, compaction and strand separation depending on the size and hence charge on the AuNPs. Moreover, this opens up to explore the exact sequence of structural changes that the DNA experience and if it is sequence specific at all. Thus the results establish an alarming message to tune and balance the charges around the nanoparticles properly before using them for different therapeutic and other application.

ASSOCIATED CONTENT Supporting Information: Details of the characterization of AuNP, TEM image, smFRET instrument setup, absorption spectrum, time-resolved data, kinetic analysis of CD measurement



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

Page 22 of 28

and individual smFRET time traces are included in SI. This information can be found on the internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author [email protected] Author Contributions #

These authors contributed equally.

Notes The authors declare no competing financial interests. ACKNOWLEDGMENT This work was funded by BARD project under the Department of Atomic Energy (DAE, Government of India). We thank, Prof Tae-Hee Lee, Pennsylvania State University for his generous help in the single molecule FRET experiments and providing the data analysis codes. We thank IIT Indore and IACS, Kolkata for providing Viscosity experiments, fluorescence intensity decay assay and the Zeta potential measurements.

REFERENCES (1) Mirkin, C. A. Programming the assembly of two- and three-dimensional architectures with DNA and nanoscale inorganic building blocks. Inorg. Chem. 2000, 39, 2258-2272. (2) Warner, M. G.; Hutchison, J. E. Linear assemblies of nanoparticles electrostatically organized on DNA scaffolds. Nat. Mater. 2003, 2, 272-277. (3) Warpehoski, M. A.; Hurley, L. H. Sequence selectivity of DNA covalent modification. Chem. Res. Toxicol. 1988, 1, 315-333.


Page 23 of 28

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


(4) Brglez, J.; Nikolov, P.; Angelin, A.; Niemeyer, C. M. Designed intercalators for modification of DNA origami surface properties. Chem. Eur. J. 2015, 21, 9440-9446. (5) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 1996, 382, 607-609. (6) He, L.; Musick, M. D.; Nicewarner, S. R.; Salinas, F. G.; Benkovic, S. J.; Natan, M. J.; Keating, C. D. Colloidal Au-enhanced surface plasmon resonance for ultrasensitive detection of DNA hybridization. J. Am. Chem. Soc. 2000, 122, 9071-9077. (7) Storhoff, J. J.; Elghanian, R.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L. One-pot colorimetric differentiation of polynucleotides with single base imperfections using gold nanoparticle probes. J. Am. Chem. Soc. 1998, 120, 1959-1964. (8) Brown, K. A.; Park, S.; Hamad-Schifferli, K. Nucleotide−surface interactions in DNAmodified Au−nanoparticle conjugates: Sequence effects on reactivity and hybridization. J. Phys. Chem. C 2008, 112, 7517-7521. (9) Tshikhudo, T. R.; Wang, Z.; Brust, M. Biocompatible gold nanoparticles. Mater. Sci. Technol. 2004, 20, 980-984. (10) Fan, J. H.; Hung, W. I.; Li, W. T.; Yeh, J. M. In 13th international conference on biomedical engineering; Lim, C., Goh, J. H., Eds.; Springer Berlin Heidelberg: 2009; Vol. 23, p 870-873. (11) Cui, P.; Seo, S.; Lee, J.; Wang, L.; Lee, E.; Min, M.; Lee, H. Nonvolatile memory device using gold nanoparticles covalently bound to reduced graphene oxide. ACS Nano 2011, 5, 68266833. (12) Homberger, M.; Simon, U. On the application potential of gold nanoparticles in nanoelectronics and biomedicine, 2010; Vol. 368.



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

(13) Tiwari, P. M.; Vig, K.; Dennis, V. A.; Singh, S. R. Functionalized gold nanoparticles and their biomedical applications. Nanomaterials 2011, 1, 31-63. (14) Chanda, N.; Kan, P.; Watkinson, L. D.; Shukla, R.; Zambre, A.; Carmack, T. L.; Engelbrecht, H.; Lever, J. R.; Katti, K.; Fent, G. M. et. al. Radioactive gold nanoparticles in cancer therapy: Therapeutic efficacy studies of GA-198 AuNP nanoconstruct in prostate tumorbearing mice. Nanomedicine 2010, 6, 201-209. (15) Cabuzu, D.; Cirja, A.; Puiu, R.; Grumezescu, A. M. Biomedical applications of gold nanoparticles. Curr. Top. Med. Chem. 2015, 15, 1605-1613. (16) Hunter, R. J.; White, L. R. Foundations of colloid science; Clarendon: Oxford, 1987. (17) Railsback, J. G.; Singh, A.; Pearce, R. C.; McKnight, T. E.; Collazo, R.; Sitar, Z.; Yingling, Y. G.; Melechko, A. V. Weakly charged cationic nanoparticles induce DNA bending and strand separation. Adv. Mater. 2012, 24, 4261-4265. (18) Han, G.; Martin, C. T.; Rotello, V. M. Stability of gold nanoparticle-bound DNA toward biological, physical, and chemical agents. Chem Biol Drug Des 2006, 67, 78-82. (19) Zhang, X.; Servos, M. R.; Liu, J. Surface science of DNA adsorption onto citrate-capped gold nanoparticles. Langmuir 2012, 28, 3896-3902. (20) Li; Rothberg, L. J. Label-free colorimetric detection of specific sequences in genomic DNA amplified by the polymerase chain reaction. J. Am. Chem. Soc. 2004, 126, 10958-10961. (21) Li, H.; Rothberg, L. Colorimetric detection of DNA sequences based on electrostatic interactions with unmodified gold nanoparticles. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 1403614039. (22) Nelson, E. M.; Rothberg, L. J. Kinetics and mechanism of single-stranded DNA adsorption onto citrate-stabilized gold nanoparticles in colloidal solution. Langmuir 2011, 27, 1770-1777.


Page 24 of 28

Page 25 of 28

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


(23) Liu, J. Adsorption of DNA onto gold nanoparticles and graphene oxide: Surface science and applications. PCCP 2012, 14, 10485-10496. (24) Zeng, S.; Yong, K.-T.; Roy, I.; Dinh, X.-Q.; Yu, X.; Luan, F. A review on functionalized gold nanoparticles for biosensing applications. Plasmonics 2011, 6, 491-506. (25) Ojea-Jiménez, I.; García-Fernández, L.; Lorenzo, J.; Puntes, V. F. Facile preparation of cationic gold nanoparticle-bioconjugates for cell penetration and nuclear targeting. ACS Nano 2012, 6, 7692-7702. (26) Mohammed, F. S.; Cole, S. R.; Kitchens, C. L. Synthesis and enhanced colloidal stability of cationic gold nanoparticles using polyethyleneimine and carbon dioxide. ACS Sustain. Chem. Eng. 2013, 1, 826-832. (27) Prado-Gotor, R.; Grueso, E. A kinetic study of the interaction of DNA with gold nanoparticles: Mechanistic aspects of the interaction. PCCP 2011, 13, 1479-1489. (28) Darbha, G. K.; Singh, A. K.; Rai, U. S.; Yu, E.; Yu, H.; Chandra Ray, P. Selective detection of mercury (ii) ion using nonlinear optical properties of gold nanoparticles. J. Am. Chem. Soc. 2008, 130, 8038-8043. (29) Liu, X.; Atwater, M.; Wang, J.; Huo, Q. Extinction coefficient of gold nanoparticles with different sizes and different capping ligands. Colloids Surf. B. Biointerfaces 2007, 58, 3-7. (30) Ha, T.; Selvin, P. R. Single-molecule techniques: A laboratory manual; Cold Spring Harbor Laboratory Press, 2008. (31) Lee, T.-H. Extracting kinetics information from single-molecule fluorescence resonance energy transfer data using hidden markov models. J. Phys. Chem. B 2009, 113, 11535-11542.



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

Page 26 of 28

(32) Boger, D. L.; Fink, B. E.; Brunette, S. R.; Tse, W. C.; Hedrick, M. P. A simple, highresolution method for establishing DNA binding affinity and sequence selectivity. J. Am. Chem. Soc. 2001, 123, 5878-5891. (33) McIntosh, C. M.; Esposito, E. A.; Boal, A. K.; Simard, J. M.; Martin, C. T.; Rotello, V. M. Inhibition of DNA transcription using cationic mixed monolayer protected gold clusters. J. Am. Chem. Soc. 2001, 123, 7626-7629. (34) Neidle, S. Nucleic acid structure and recognition; Oxford University Press: Oxford, U.K., 2002. (35) Grueso, E.; Perez-Tejeda, P.; Prado-Gotor, R.; Cerrillos, C. DNA strand elongation induced by small gold nanoparticles at high ethanol content. J. Phys. Chem. C 2014, 118, 4416-4428. (36) Rosenberg, A. H.; Studier, F. W. Intrinsic viscosity of native and single-stranded T7 DNA and its relationship to sedimentation coefficient. Biopolymers 1969, 7, 765-774. (37) Ray, P. C.; Fortner, A.; Darbha, G. K. Gold nanoparticle based fret assay for the detection of DNA cleavage. J. Phys. Chem. B 2006, 110, 20745-20748. (38) Eggeling, C.; Fries, J. R.; Brand, L.; Gunther, R.; Seidel, C. A. Monitoring conformational dynamics of a single molecule by selective fluorescence spectroscopy. Proc Natl Acad Sci U S A 1998, 95, 1556-1561. (39) Widengren, J.; Schwille, P. Characterization of photoinduced isomerization and backisomerization of the cyanine dye cy5 by fluorescence correlation spectroscopy. J. Phys. Chem. A 2000, 104, 6416-6428. (40) Tinnefeld, P.; Buschmann, V.; Weston, K. D.; Sauer, M. Direct observation of collective blinking and energy transfer in a bichromophoric system. J. Phys. Chem. A 2003, 107, 323-327.


Page 27 of 28

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


(41) Hofkens, J.; Cotlet, M.; Vosch, T.; Tinnefeld, P.; Weston, K. D.; Ego, C.; Grimsdale, A.; Müllen, K.; Beljonne, D.; Brédas, J. L. Revealing competitive Förster-type resonance energytransfer pathways in single bichromophoric molecules. Proc. Natl. Acad. Sci. 2003, 100, 1314613151. (42) Deo, S.; Godwin, H. A. A selective, ratiometric fluorescent sensor for Pb2+. J. Am. Chem. Soc. 2000, 122, 174-175. (43) Mello, J. V.; Finney, N. S. Dual‐signaling fluorescent chemosensors based on conformational restriction and induced charge transfer. Angew. Chem. 2001, 113, 1584-1586.



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

TOC


Page 28 of 28

Conformational Changes Followed by Complete Unzipping of DNA

Recommend Documents