Fluorimetric detection of G-quadruplex DNA in solution and adsorbed

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Fluorimetric detection of G-quadruplex DNA in solution and adsorbed on surfaces with a selective trinuclear cyanine dye Heiko Ihmels, Siyu Jiang, Mohamed Mahmoud, Holger Schönherr, Daniel Wesner, and Imad Zamrik Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b02382 • Publication Date (Web): 01 Sep 2018 Downloaded from http://pubs.acs.org on September 3, 2018

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Fluorimetric detection of G-quadruplex DNA in solution and adsorbed on surfaces with a selective trinuclear cyanine dye Heiko Ihmels,*Siyu Jiang, Mohamed M. A. Mahmoud, Holger Schönherr,* Daniel Wesner, Imad Zamrik ‡ Department of Chemistry and Biology, University of Siegen, and Center of Micro- and Nanochemistry and Engineering (Cµ); Adolf-Reichwein-Str. 2, 57068 Siegen.

‡

Author names are in alphabetical order that does not reflect the specific contribution of each author.

E-mail: [email protected]; [email protected]

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Abstract

Quadruplex DNA, which is a relevant target for anticancer therapies, may alter its conformation due to interactions with interfaces. In pursuit of a versatile methodology to probe adsorption-induced conformational changes, the interaction between a fluorescent [2.2.2]heptamethinecyanine dye and quadruplex DNA (G4-DNA) was studied in solution and on surfaces. In solution, the cyanine dye exhibits a strong light up effect upon the association with G4-DNA without interference from double stranded DNA. In addition, a terminal πstacking as a binding mode between the cyanine dye and G4-DNA is concluded using NMR spectroscopy. To unravel the effects of adsorption on the conformation of quadruplex-DNA, G4-DNA, double stranded and single stranded DNA were adsorbed to positively charged poly(allylamine) (PAH) surfaces, both in planar and in constrained 55 nm diameter aluminum oxide nanopore formats. All DNA forms showed a very strong affinity to the PAH surfaces as shown by surface plasmon resonance and reflectometric interference spectroscopy, respectively. The significant increase of the fluorescence emission intensity of the cyanine light up probe observed exclusively for surface immobilized G4-DNA affords evidence for the adsorption of G4-DNA on PAH with retained quadruplex conformation.


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Introduction The telomere is a DNA-protein complex at the end of the chromosome that has essential biological functions.1,2,3 It contains a single-stranded guanine-rich oligonucleotide overhang with a repetitive d(TTAGGG) sequence, the so called telomeric DNA, that protects the chromosomes from enzymatic degradation.4,5 In normal cells, the telomeric DNA overhang becomes shorter with each cell cycle leading to senescence and eventually cell death; however, in cancer cells the enzyme telomerase maintains the telomeric DNA length, so that these cells become potentially immortal.6 Therefore, telomerases are a promising key target in cancer therapy.7,8,9,10 In this context, it has been shown that G-rich sequences have the ability to fold into G-quadruplex (G4-DNA) structures,11,12,13,14,15,16,17 and it has been proposed that the formation and stabilization of quadruplex structures in the telomeric DNA may inhibit the telomerase activity because it interferes with the formation of the telomeretelomerase complex.12,13,18,19 Therefore, designing small molecules that stabilize G4-DNA at the telomeric end is considered as a useful strategy to develop anticancer therapeutic agents.18,20,21 During the past decades, several organic and organometallic molecules have been developed to selectively associate with G4-DNA.4,12,20,22,23,24,25,26,27 In addition, the development of efficient fluorescent probes for the selective detection of G4-DNA has great potential due to the high sensitivity of fluorescence spectroscopy.28,29,30,31 From the already reported G4-DNA ligands, cyanine dyes are attractive representatives as they show fluorescence light-up effects upon association with the G4-DNA.32,33 Among these cyanine dyes, the trinuclear cyanine dyes 1a and 1b (Chart 1) show a moderate binding affinity with the telomeric G4-DNA (Kb = 8.5−9.5 × 105 M-1) and light-up a factor of 106 for 1a and 64 for 1b upon association with G4-DNA with high selectivity in comparison to the double-stranded DNA.34,35 Recently, the heptamethinecyanine dye 1c was also introduced as a promising G4DNA ligand that exhibits a fluorescence light up factor of 190 upon association with


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telomeric quadruplex DNA, which is the most effective fluorimetric response in the series of studied trinuclear cyanine dyes.36 However, no selectivity studies were performed thus far to assess the ability of cyanine dye 1c to differentiate between different types of DNA. Similar to the study of DNA-binding light up probes, the adsorption or immobilization of DNA on surfaces is a fundamental research area. It also possesses relevance for developing chip-based DNA sensors, for fabrication of DNA nanostructures and for gene delivery techniques.37,38,39,40,41,42 One of the promising methods to adsorb DNA on surfaces is inspired by the layer-by-layer (LbL) technique, which is based on the alternating deposition of layers of anionic and cationic polymers.43,44,45 In addition, using the LbL assembly method in 3Dtemplates such as nanoporous anodic aluminum oxide (AAO) is considered an attractive tool to prepare biomacromolecule nanotubes and optical biosensors.46,47,48 However, thus far G4DNA biosensors were prepared, to our knowledge, only by using a linker such as biotin to interact with a streptavidin-coated surface.49,50,51 In this work, we analyzed the binding of various forms of DNA on positively charged poly(allyl amine) (PAH) modified planar surfaces and curved nanoporous substrates. In particular, the interaction between the representative cyanine dye 1c, which holds ideal features of a DNA-sensitive fluorescent light-up probe, and DNA immobilized on the planar surfaces and inside geometrically constrained nanopores was investigated to unravel whether or not G4-DNA adsorbed in an intact manner. The polyelectrolyte modified AAO nanopores allow for the variation of the curvature and interactions so that interfaces e.g. within the cell or the nucleus can be modeled using this system.


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Chart 1. Structures of cyanine dyes 1a− −c.

Results and Discussion Thermal DNA-denaturation experiments In order to assess the stabilization effect of the cyanine dye 1c toward G4-DNA, the melting temperature, Tm, of the dye-labelled quadruplex-forming oligonucleotide F21T [fluorescein-d(G3T2AG3T2AG3T2AG3)-tetramethylrhodamine] was determined. In the folded quadruplex structure, both dyes are located close enough to ensure a Förster resonance energy transfer (FRET) from the excited fluorescein to the rhodamine unit,52 whereas in the unfolded single strand the FRET is no longer possible due to the large separation of the dyes. Thus, the melting curve and the melting temperature Tm, are readily determined by a plot of the emission intensity at a given wavelength, as an indicator of the FRET efficiency, versus temperature. The ligand-induced stabilization of the G4-DNA towards unfolding is then quantified by the shift of the melting temperature, ∆Tm, in the presence of the ligand.53 The melting temperature of the DNA F21T is Tm = 45.5 °C (Figure 1).54 Upon the addition of the cyanine dye 1c, the melting temperature increased strongly by up to ∆Tm = 25.5 °C at a ligand-DNA ratio (LDR) of 5 (Figure 1). To study the selectivity of the cyanine dye 1c with regard to differentiation between duplex and quadruplex DNA, the melting temperatures of the quadruplex F21T at different LDR values were determined in the presence of an excess (15 molar equiv. with respect to the G4-DNA) of the self-complementary oligonucleotide


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ds26 [d(CA2TCG2ATCGA2T2CGATC2GAT2G]. In general, the ligand-induced melting temperature of the quadruplex DNA is only slightly lower (∆∆Tm ≈ 3 °C at LDR = 2.5 and 5.0; Figure 1) in the presence of the duplex DNA ds26.

Figure 1. Fluorimetric analysis of thermal denaturation of G4-DNA F21T (0.2 µM) in the presence of cyanine 1c; LDR: 0, 1.3, 2.5, 5 (molar equivalents); in KCl-LiCl-Na-cacodylate buffer (10 mM K+, 10 mM Na+, 90 mM Li+, pH 7.2; λex = 470 nm; λem = 515 nm). The arrows indicate the evolution of the melting curves with increasing LDR. Inset: Induced shift of melting temperature (∆Tm) of F21T upon the addition of cyanine 1c in absence () and presence (○) of ds26. Photometric and fluorimetric titrations The

interaction

of

the

cyanine

1c

with

telomeric

G4-DNA

22AG

[d(AG3T2AG3T2AG3T2AG3)] was monitored by photometric and fluorimetric titrations (Figure 2A). The dye 1c exhibits a broad long-wavelength absorption maximum at 626 nm. Upon addition of 22AG, a hypochromic effect was observed at LDR > 7. Further addition of DNA resulted in a bathochromic shift (∆λ = 24 nm) with a hyperchromic effect. At the same time, the addition of double-stranded calf thymus (ct) DNA resulted only a small light-up factor of only 20. The analysis of the resulting binding isotherm revealed a binding constant


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between 1c and ct DNA of Kb = 4.0 ×105 M–1 (cf. SI: Figure S1). Based on these different effects of the DNA on the emission properties of the cyanine 1c, the selectivity of the ligand 1c towards G-quadruplex and duplex DNA was further examined by fluorimetric detection of a competition experiment. Hence, a mixture of cyanine 1c and ct DNA (LDR = 0.1) was titrated with a solution of G4-DNA 22AG (Figure 2B). The weak emission of the cyanine 1c when bound to ct DNA increased drastically with the addition of 22AG. In an inverse control experiment, the titration of ct DNA to a solution of cyanine 1c and G4-DNA 22AG (LDR = 0.1) did not result a significant change of the fluorescence intensity of the ligand.

Figure 2. A: Photometric titration of 1c (c1c = 10 µM) with G4-DNA 22AG. Arrows indicate the development of the absorbance with increasing DNA concentration. B: Plot of the relative fluorescence intensity of 1c (c1c = 2.5 µM) versus molar fraction X22AG as obtained from the addition of 22AG to a mixture of ct DNA (25 µM) with 1c (), or the addition of ct DNA to a mixture of 22AG (25 µM) with 1c (); in potassium phosphate buffer (95 mM, pH 7.0);

λex = 580 nm.

The continuous variation method (Job plot analysis)55 was applied to determine the binding stoichiometry between the cyanine 1c and G4-DNA 22AG. The emission intensity of


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mixtures of the ligand 1c were determined at different fractional ratios c1c : c22AG (with constant total concentration). The data were presented as a Job plot and the intercept of the straight lines, as resulting from the linear regression of ascending and descending data points, respectively, was located at XLigand = 0.65 which corresponds to a binding stoichiometry of essentially 2 ligands per G4-DNA (Figure 3).

Figure 3. Job plot obtained from the fluorimetric analysis of solutions with different ratios of 1c and 22AG (c1c + cDNA = 10 µM) in potassium phosphate buffer (95 mM, pH 7.0); X2b = mole fraction of ligand; λex = 580 nm.

1

H-NMR spectroscopy

To acquire information on the binding site that is possibly occupied by the cyanine 1c, the 1

H-NMR spectra of the complex between Tel26 [d(A3G3T2AG3T2AG3T2AG3A2)] (Figure 4)

and ligand 1c was investigated. In our hand, the 1H-NMR spectra of the Tel26 show only slight deviation from the reported data which does not affect further analysis of ligand-DNA complex.56,57 The addition of cyanine 1c leads only to slight but significant changes of the chemical shifts of several imino and aromatic protons in the 1H-NMR spectrum of the quadruplex DNA Tel26 (Figure 5 and 6). Specifically, the imino protons of the G10, G11, G17, G22 and G23 (cf. SI: Figure S2) as well as the aromatic protons of G4, G10, G17, T19, A21, G22 and G23 show a small, but significant shift by 0.02 ppm. During the titration, the


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proton signals of the cyanine 1c were not detected which is often observed in G4-DNAligand complexes.58,59

Figure 4. Schematic representation of the folding topology of G4-DNA Tel26 in K+containing buffer; red: deoxyguanosine (blue indicates syn conformation); green: deoxyadenosine; black: deoxythymidine.


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Figure 5. 1H-NMR spectra (600 MHz) of Tel26 (2.0 mM in bases) in the range of the imino protons (10.5−12.1 ppm) with increasing amount of cyanine 1c; in K-phosphate buffer (95 mM, H2O : D2O = 9 : 1; pH 7.0); T = 31 °C.

Figure 6. 1H-NMR spectra (600 MHz) of Tel26 (2.0 mM in bases) in the range of the aromatic protons (6.6−8.3 ppm) with increasing amount of cyanine 1c; in K-phosphate buffer (95 mM, H2O : D2O = 9 : 1; pH 7.0); T = 31 °C. To further monitor the interaction of the cyanine 1c with Tel26, NOESY-NMR experiments were performed (Figure 7). Considering the higher temperature used in our experiment than the reported one (31 °C instead of 1 °C), some of the reported cross peaks were not detected, however, most of the characteristic intra- and interquartet NOE cross peaks of Tel26 were observed, such as G6H1-G16H1, G5H1-G6H1, G11H1-G12H1, G11H1-G18H1 and G23H1-G24H1.56 The addition of the cyanine 1c induces slight changes in some NOE cross peaks of Tel26. Particularly, the cross peaks of G10-G4, G10-G5 and G10-G18 in the imino region, which relates to the shifts of the corresponding 1H-NMR


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signals, show the most significant changes (Figure 7). In addition, the cross peaks of G10H1'G10H8, T19H1'-T19H6, T20H1'-T20H6 and A12H1'-A21H6 show noticeable shift in the presence of the cyanine 1c (Figure 7).

Figure 7. Superposed 2D NMR spectra (NOSEY) of Tel26 (2.0 mM, in bases) in the absence (blue) and in the presence (red) of cyanine 1c (0.8 molar equivalent); H2O-D2O (9:1); Kphosphate buffer (95 mM, pH 7.0); T = 31 °C. ACS Paragon Plus Environment

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DNA deposition on planar surface and inside nanopores The deposition of G4-DNA 22AG, double stranded DNA ds26 and single stranded DNA ss26, [d(CA2TCG2ATCGA2CA2TCG2ATCGA2)] on positively charged poly(allylamine) hydrochloride (PAH) layers was investigated on gold-coated planar surfaces and inside anodic aluminum oxide (AAO) nanopores. Firstly, the adsorption and desorption behavior of G4-DNA 22AG on PAH-coated planar surfaces was determined using surface plasmon resonance (SPR) measurements (Figure 8).

Figure 8. SPR measurements of adsorption and desorption of G4-DNA 22AG on PAHcoated planar samples (c22AG = 10 µM; K-phosphate buffer: 95 mM, pH 7.0). (A) Kinetic SPR data of adsorption of G4-DNA 22AG acquired at fixed angle. (B) SPR reflectivity scans taken before (black curve) and after (red curve) adsorption of G4-DNA 22AG. (C) SPR ACS Paragon Plus Environment

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reflectivity scans taken after buffer rinse (black curve) and after rinsing continuously with pure buffer for 12 h (red curve). The addition of 22AG to PAH-coated surfaces resulted in a significant shift of the minimum of the curve of the angular SPR reflectivity scan with ∆θ = 520 m° (Figure 8B). After solution exchange the subsequent continuous pumping of K-phosphate buffer through the SPR cell for more than 12 h led to a minute decrease of the SPR angle shift of only 18 m° (Figure 8D), which indicates the very strong, irreversible binding of G4-DNA to PAH-coated surfaces. To further study the adsorption kinetics of G4-DNA 22AG on PAH-coated planar surfaces and inside the nanopores, the effect of different concentrations of 22AG was studied (Figure 9 and Table 1). While the adsorption of G4-DNA on planar surfaces was followed by SPR, the kinetics inside AAO was followed in situ by reflectometric interference spectroscopy (RIfS).60 In both cases, the apparent observed rate constants (kobs) increased with increasing concentration. However, inside the AAO nanopores the constants were generally significantly lower. Notably, the association rate constant (kon) of the G4-DNA on planar surfaces was one magnitude higher than that determined inside the AAO nanopores (kon = 4.2 × 10–3 ± 1.2 × 10– 4

M–1 s–1 on the planar surface and 5.5 × 10–4 ± 1.5 × 10–5 M–1 s–1 inside the AAO nanopores).

Likewise, the dissociation rate constant (koff) of the G4-DNA on planar surfaces was one magnitude higher than that determined inside the AAO nanopores (koff = 9.7 × 10–4 ± 1.3 × 10–5 s–1 on planar surface and 5.4 × 10–5 ± 1.7 × 10–6 s–1 inside the AAO nanopores). Due to the well-known limited sensitivity of SPR in detecting the adsorption of minute amounts of low molar mass compounds to the sensor surface,61 the interaction between cyanine 1c and surface immobilized G4-DNA was not detected by SPR (no data shown). Likewise, also by RIfS no change in the effective optical thickness was observed upon the addition of 1c to DNA covered AAO nanopores.


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Table 1. Observed apparent rate constants (kobs) for the adsorption of the G4-DNA 22AG on PAH-coated surfaces, as obtained from SPR and RIfS. Planar surface

AAO Nanopores

c22AG /µM

kobs /10–3 s–1

kobs/ 10–3 s–1

0.5

3.3

0.5

1.0

4.5

0.7

5.0

20

2.5

10

46

5.3

20

83

11

a

b

a

Observed rate constants (kobs) for adsorption of G4-DNA 22AG on PAH-coated planar surfaces. b Observed rate constants (kobs) for adsorption of G4-DNA 22AG on inside AAO nanopores coated with PAH. All values were calculated from fits of the experimental SPR and RIfS data using eq. 2 (cf. Experimental Section).

Figure 9. Kinetics of the adsorption of G4-DNA 22AG on PAH-coated surfaces. (A) SPR data for planar substrates. (B) RIfS data for AAO nanopores. Insets: Plots of kobs versus DNA concentration; the red lines denote the best fit of the experimental data using equation 2 or equation 4 (cf. Experimental Section).


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From the SPR and RIfS experiments carried out at different DNA concentrations, the adsorption isotherms of G4-DNA 22AG, ss-DNA ss26 and ds-DNA ds26 were determined. The data are characteristic of high affinity isotherms, consistent with the very strong attachment of DNA on PAH surfaces (Figure 10 and Table 2). The maximum surface excess (Γmax) observed on both planar surfaces and inside the nanopores indicates that the mass coverage of the 22AG is higher than the one of ss26 than ds26 (Table 2). Remarkably, an apparent saturation of the PAH-coated surface was achieved at 5.0 µM for all types of DNA for both the planar surfaces or inside the AAO nanopores (Figure 10). Therefore, the thickness of DNA adsorbed at a concentration of 5.0 µM on the planar surfaces was estimated to be 2.0, 1.6 and 1.4 nm for 22AG, ss26 and ds26, respectively. By contrast, inside the AAO nanopores the thickness was estimated to be only 0.91 nm, 0.80 nm and 0.34 nm for 22AG, ss26 and ds26, respectively. The thicknesses were calculated according to the volume fractions concluded from equation 9 (cf. Experimental Section).

Table 2. Maximum surface excess (Γmax), Langmuir constant (KL) and layer thickness (l) of DNA on the PAH-coated planar surfaces and inside the AAO nanopores. DNA

Planar surface

Γmax / m°

KL / µM–1

a

b

22AG

525

2.6

ss26

427

ds26

360

AAO nanopores

Γmax / nm

KL / µM–1

l / nm f

d

e

2.0

14.8

1.8

0.91

2.4

1.6

13

1.6

0.80

2.2

1.4

5.6

1.5

0.34

l / nm c

a

Maximum surface excess determined by SPR using eq. 5. b Langmuir constant determined from the SPR data using eq. 5. c Thickness, l, determined by SPR using Winspall software. d Maximum surface excess determined from the RIfS data using eq. 5. e Langmuir constant determined by RIfS using eq. 5. f Thickness determined by RIfS using eq. 8.


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Figure 10. Adsorption isotherms of G4-DNA 22AG (black), ss26 (red) and ds26 (blue) on PAH-coated planar surfaces as obtained (A) from the SPR and (B) the RIfS experiments (Kphosphate buffer: 95 mM, pH 7.0). The solid lines denote the best fits of the experimental data using eq.5. Confocal fluorescence microscopy (CFM) Since SPR and RIfS are not sensitive enough to detect the interaction of cyanine 1c with immobilized DNA on PAH-coated surfaces, confocal fluorescence microscopy (CFM) experiments were employed (Figure 11; cf. SI: Figure S3 and S4). The deposited 22AG and ss26 on the PAH-coated planar surfaces and inside the PAH-coated AAO nanopores did not show marked fluorescence upon excitation with a red laser (λ = 635 nm). By contrast, the addition of cyanine 1c onto G4-DNA 22AG adsorbed on a planar surface resulted in a significant increase in the fluorescence intensity, which was more than 160 times stronger than for ss26 modified surfaces (Figure 11A and 11B). Likewise, the fluorescence intensity of cyanine 1c upon binding with G4-DNA 22AG inside the AAO nanopores was 32 times higher than for ss26 (Figure 11C and 11D). Notably, the background fluorescence intensity observed for cyanine 1c inside the PAH covered AAO nanopores was significantly higher than that observed on planar surface.


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Figure 11. Fluorescence intensity distributions obtained by CFM on PAH-coated surfaces in the absence of ligand 1c (black columns) and in the presence of 1c (green columns). (A, B) Planar surfaces and (C, D) AAO nanopores with immobilized 22AG (A and C) and ss26 (B and D) (cDNA = 5.0 µM; c1c = 2.5 µM); K-phosphate buffer: 95 mM, pH 7.0). The solid red lines denote fits of the experimental data by a normal distribution affording mean values of 347 and 10 for 22AG and ss26, respectively. By contrast, the mean values for planar substrates were 167 and 2 for 22AG and ss26, respectively.

To analyze the binding kinetics of G4-DNA and the dye 1c in solution also a confocal fluorescence microscopy setup was used, since the kinetics was slow enough to be followed, when working in the nM range (Figure 12A). The low initial fluorescence intensity of the free


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cyanine dye dropped due to the dilution, when the DNA solution was added, and then immediately started to rise, indicating the binding to the G4-DNA. The fitted exponential curves afforded apparent rate constants kobs, which depend on the concentration of the DNA (eq.4). From the linear fit (Figure 12B) values of the rate constants kon = 2.7 × 105 ± 0.4 × 105 M-1 s-1 and koff = 2.9 × 10-2 ± 1.4 × 10-2 s-1 were derived. The scatter for the data for different DNA concentrations is attributed to the manual mixing of the reaction solution of 20 µL by hand using a pipette. The ratio of kon and koff yields a binding constant Kb = 9.3 × 106 M-1.

Figure 12. (A) Temporal evolution of fluorescence intensity of a solution containing the dye 1c before and after addition of G4-DNA, as measured by CFM (initial dye concentration 200 nM, after addition of DNA solution diluted to 20 nM). The fluorescence intensity of the free dye 1c increases upon binding to G4-DNA following an exponential curve (eq. 3, red line). (B) Analysis of kinetic data according to eq. 4. The observed apparent rate constants kobs depend linearly on the DNA concentration, yielding the rate constants kon and koff.


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Discussion Binding interactions between 1c and DNA in solution All the experimental results point to a selective association of the ligand 1c with quadruplex DNA in solution. Firstly, the change of the absorption spectrum of cyanine 1c upon the addition of G4-DNA 22AG indicates that 1c associates with the telomeric quadruplex. It is proposed that the absorption of the free dye 1c with a maximum at 626 nm is due to the formation of aggregates in aqueous solution as commonly observed for such cyanine dyes.62,63 Thus, the bathochromic shift that is observed by the addition of the DNA along with an increase of the intensity is induced by the disassembly of the cyanine 1c aggregates and binding of the monomer to the G4-DNA.34,64 The lack of isosbestic points during titration (Figure 2) indicates different binding modes between 1c and G4-DNA that obviously depend on the LDR value, i.e. on the availability of binding sites. The considerable increase in the melting temperature of the dye labeled G4-DNA F21T upon the association with cyanine 1c (∆Tm = 25.5 °C at LDR = 5) indicates a relatively strong stabilization of G4DNA on complexation of 1c, as compared e.g. with the resembling cyanine derivative 1a (∆Tm = 8.5 °C at LDR = 5).34 This observation is in agreement with the favorable stabilizing effect of quinolinium fragments of quadruplex ligands as this type of cationic heterocycle is frequently used in high-affinity quadruplex ligands.65,66 Most notably, the addition of doublestranded DNA ds26 does not affect the stabilization effect of cyanine 1c on F21T significantly (∆∆Tm = 3.3 °C), which indicates high selectivity of cyanine 1c towards Gquadruplex as compared to double-stranded DNA. The binding of the cyanine 1c with G4DNA is further confirmed by NMR-spectroscopy. The NMR-spectra of the Tel26-cyanine 1c complex show a possible terminal π-stacking binding mode at the G4-G10-G18-G22 quartet as the 1H-NMR signals of the guanine bases of the terminal quartet G4, G10 and G22 as well as the signals of the loop residue T19, T20 and A21 were the most affected upon the


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association with the cyanine 1c. These assignments were supported by the NOE-NMR data of the ligand-DNA complex as the shifts are more noticeable in the 2D presentation of the spectra (Figure 7). Specifically, the G4-G10-G18-G22 NOE cross peaks as well as the ones between the sugar residues and the aromatic protons of the loop residue T19, T20 and A21 are indicative of the proposed binding mode. Moreover, the intercalation binding mode between 1c and Tel26 is excluded as in the presence of the cyanine 1c none of the NOE cross peaks between the imino protons of the guanine bases disappeared. It should be noted that the shift difference is only very small, but still distinctly larger than the experimentally determined standard deviation ± 0.002 ppm of the chemical shift. In general, cyanine dyes exhibit low emission quantum yields due to the radiationless deactivation of the excited state mainly caused by conformational relaxation or E-Z isomerization.34,67 It is well known that the accommodation of these dyes in the binding sites of host molecules such as nucleic acids suppresses these deactivation pathways thus leading to strong increase of the emission quantum yield.68 Accordingly, the very weak fluorescence intensity of the cyanine 1c increases significantly upon association with G4-DNA by a factor of about 190.36 Although such light-up factors are only an approximation because the starting intensity is very small, a comparison shows that this value is significantly higher than the light-up effect of resembling cyanine derivatives 1a (factor: 106) and 1b (64) under similar conditions.34,36 Most notably, the light-up effect of 1c is more pronounced with quadruplex DNA than with duplex DNA, and this difference could be used to assess the selectivity of the ligand towards G4-DNA in a competition experiment (Figure 2). Especially the light-up effect caused by the addition of the G4-DNA to a complex of ct DNA and cyanine 1c indicates that the ligand is displaced from the double-stranded DNA and redistributed into Gquadruplex binding sites, which reflects the higher affinity of 1c towards G4-DNA with a binding constant of 0.9 x 106 M−1 between 1c and G4-DNA.36


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The analysis of the binding kinetics in solution makes use of the increase of the fluorescence intensity of 1c upon binding to G4-DNA. The high sensitivity of the confocal detection system allowed for working in the nM concentration range so that the otherwise fast binding kinetics could be measured. Nevertheless; the scattered data are due to the relatively low fluorescence intensity at this concentration. The total intensity increase might not reach the maximum value given above, since not all free dye molecules will bind to DNA at a concentration that is of the same order of magnitude as the dissociation constant determined to be KD = 1/Kb = 110 nM. The KD value reported before36 this is one order of magnitude larger so that this effect might be even more pronounced.

Binding interactions of quadruplex DNA and DNA-ligand complexes with surfaces The results obtained from the SPR point to a very strong binding of the G4-DNA 22AG to the positively charged PAH covered surface with a negligible koff value (10-5 s-1). Therefore, the adsorption of the G4-DNA on PAH covered surface is considered to be irreversible. Together with the kinetic parameters for the binding reaction in solution a more sophisticated picture of the interactions involved in the binding process of quadruplex DNA and the ligand 1c on surfaces was sought (cf. SI: Figure S6). The irreversible adsorption of the G4-DNA helped to simplify this model. Notably, the adsorption of G4-DNA 22AG is 8 times faster on the PAH covered planar surface than inside the PAH covered AAO nanopores. The slow adsorption of the G4-DNA inside the nanopores is attributed to the mass transport into these pores. It has been reported that nanopores function as reacting surface (perfect sink conditions), which produces a stationary concentration profile.69 Due to the confined space within the nanopores the DNA adsorption might be affected by the electrostatic charge of the neighboring DNA molecules already bound on the surface. This can explain the reduced average thickness of the adsorbed layer as well as the somewhat smaller Langmuir constant


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in the nanopores compared to the flat surface. In addition, G4-DNA tends to deposit as large and ordered spherical aggregates leading to a higher surface coverage than ss-DNA and dsDNA.70 On the other hand, ss-DNA exhibits higher hydrophobicity, lower linear charge density and lower persistence length than ds-DNA, which leads to higher surface excess of ss-DNA compared to ds-DNA and concomitantly higher mass coverage.71,72,73 The addition of the cyanine 1c to the surface-immobilized G4-DNA 22AG on planar surfaces produced a significantly higher increase in emission intensity than that observed with the immobilized ss26. This increase exceeded the increase in mass coverage by a factor of 168, hence providing evidence that 22AG preserved to some significant extent its quadruplex topology on the surface during adsorption. The absolute emission intensity of the cyanine 1c was higher inside the AAO nanopores than on planar surface. This observation is attributed to the 3-dimensional sample geometry with cylindrical pores, which results in a larger surface area and consequently to an increased number of DNA-bound ligands within the same confocal volume compared to the planar 2-dimensional samples. From the data acquired no evidence for the impact of curvature on the DNA conformation was obtained, which is not surprising considering the comparatively large radius of curvature of ~ 27 nm.

Conclusion All experimental results point to a selective association of the trinuclear cyanine dye 1c with G4-DNA in solution and G4-DNA adsorbed on surfaces. In solution, cyanine 1c exhibits high selectivity towards G4-DNA over double stranded DNA, as was concluded from the thermal DNA denaturation experiments and the fluorimetric titrations. In addition, the fluorescence intensity of 1c increases dramatically upon association with G4-DNA. The NMR experiments indicate a possible terminal π-stacking binding mode between 1c and G4DNA. The kinetics of this interaction has been analyzed making use of the change in


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fluorescence intensity. Furthermore G4-DNA shows an irreversible adsorption on PAHcoated polycationic surfaces. Thermodynamic and kinetic parameters of this interaction were determined for planar and geometrically constrained surfaces. Despite this strong interaction the quadruplex structure remains intact, as concluded from the binding of 1c to the immobilized G4-DNA. Since quadruplex structures interfere with telomerase activity, the stabilization of the G4-DNA using ligands like 1c can also have implications for cancer therapy.


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Experimental Materials The cyanine dye 1c was synthesized and characterized according to the published procedure.74

Oligodeoxyribonucleotides

(HPLC

purified)

Tel26

[d(A3G3T2AG3T2AG3T2AG3A2)], 22AG [d(AG3TTAG3TTAG3TTAG3)], F21T [fluoresceind(G3T2AG3T2AG3T2AG3)-tetramethylrhodamine], ss26 [d(CA2TCG2ATCGA2CA2TCG2ATCGA2)] and ds26 [d(CA2TCG2ATCGA2T2CGATC2GAT2G] were purchased from Metabion Int. AG (Planegg/Martinsried). The nucleic acids were dissolved in the corresponding buffer, heated to 95 °C for 5 min and subsequently cooled slowly over 4 h to room temperature. Double-stranded ct DNA was purchased from Sigma-Aldrich and dissolved in BPE buffer and stored at 4 °C. The concentration was determined photometrically (λmax= 260 nm, ε = 12824 cm−1 M−1).75 Potassium phosphate buffer: 25 mM K2HPO4, 70 mM KCl; adjusted with 25 mM KH2PO4 to pH 7.0; sodium cacodylate buffer: 10 mM Na(CH3)2AsO2·3H2O, 10 mM KCl, 90 mM LiCl; pH 7.2–7.3; BPE buffer: 6.0 mM Na2HPO4, 2.0 mM NaH2PO4, 1.0 mM Na2EDTA; pH 7.0. Poly(allylamine) hydrochloride PAH (Mw = 120 000 - 200 000 g/mol, Alfa Aesar), poly(sodium 4-styrenesulfonate) PSS (Mw =

70

000

g/mol,

Sigma-Aldrich),

KCl

(Sigma-Aldrich),

ethanol

(VWR),

16-

mercaptohexadecanoic acid (MHDA, 90%, Sigma-Aldrich), phosphoric acid (85%, Chemische Fabrik Budenheim), 3-(ethoxydimethylsilyl)-propylamine (97%, C.N. 18306-791, Sigma-Aldrich). Equipment Absorption spectroscopy: Varian Cary 100 Bio spectrophotometer; emission spectroscopy: Varian Cary Eclipse; NMR spectroscopy: Varian VNMR-S 600 spectrometer (1H: 600 MHz). SPR glass prisms: SF15 (n= 1.69425) Bernhard Halle Nachfl. GmbH (Berlin, Germany). Confocal fluorescence microscope: Microtime 200, PicoQuant (Berlin, Germany)


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with inverted laser scanning microscope (IX-71, Olympus, Hamburg, Germany) and a 40× objective LD Achroplan, NA 0.60 (Zeiss, Oberkochen, Germany). Interferometric measurements were performed utilizing a CCD detector (USB 2000+) equipped with a tungsten halogen light source (LS-1), and a bifurcated optical fiber (R400-7-VIS/NIR) to transfer the light to the optical probe. The reflected light was collected by the same probe and transferred to the CCD detector. All of the RIfS setup parts were purchased from Jaz, Ocean Optics, Inc., Dunedin, FL, USA. The data were captured by Spectra Suite Spectroscopy software (Jaz, Ocean Optics, Inc., Dunedin, FL, USA) in which the number of scans to average and the integration time were set to 500 scan/s and 10 s, respectively. For data analysis, fast Fourier transform (FFT) was applied (utilizing Wavemeteric IGOR Pro 6 software) and Fringes 22.6 data analysis program (obtained from Professor Michael J. Sailor, University of California, San Diego, USA) on the reflectance spectrum, which resulted in a single peak, whose position along the x-axis corresponds to the EOT value of the thin film. Scanning Electron Microscopy (SEM) Measurements: The SEM data were acquired on a Zeiss Ultra 55cv field emission scanning electron microscope (SEM) (Zeiss, Oberkochen, Germany). All measurements were performed with an operation voltage of 10 kV with the Inlens secondary electron detector. 10 nm gold was sputtered on the samples to obtain a conductive surface. For the analysis of the SEM micrographs, SPIP software (Version 5.0.7) and ImageJ software (version 1.47) were used. Methods All photometric and fluorimetric titrations were performed in thermostated quartz cuvettes at 20 °C. Titrant solutions were freshly prepared by dilution of aliquots from the stock solution. Spectrophotometer slit widths were 2 nm for photometric experiments and 5 nm for fluorimetric experiments. The binding constants were determined by fitting the binding


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isotherms from the fluorimetric titrations to the established theoretical model according to the independent-site model (eq. 1).76

I =

(eq. 1)

where n is the number of the binding sites per quadruplex DNA. Thermal DNA denaturation experiments were performed according to published procedures.34 For Job plot analysis (continuous variations method), a series of samples was prepared with a constant sum of concentrations at 10.0 µM, but with varying concentrations of ligand and DNA. The fluorescence spectra were recorded with λex = 580 nm. 1

H-NMR spectra were recorded on a Varian VNMR-S 600 spectrometer (1H: 600 MHz)

with a triple resonance HCN inverse probehead. 1D NMR spectra were recorded with solvent suppression with the pulse sequence WET1d with 1.5 s relaxation time and 256 scans. Solvent suppression in 2D NMR measurements was accomplished with the sequence WET NOESY, with mixing time of 300 ms, relaxation time of 1.5 s, and detection of 256 × 64 or 512 × 96 data points. Chemical shifts of 1H-NMR spectra are given in ppm (δ) relative to 4,4dimethyl-4-silapentane-1-sulfonic acid (DSS, δ = 0.00 ppm). Highly ordered AAO nanopores were fabricated using a two-step electrochemical anodization process,77 followed by chemical widening by immersing in 0.5 M phosphoric acid solution at 33°C for 15 min. The resulting AAO pores were characterized by SEM (cf. SI, Figure S5). The anodization condition employed here produced pore diameters of 55 ± 5 nm, interpore distances of 112 ± 2 nm, and wall thicknesses of 75 ± 2 nm. The calculated porosity and pore density were 22% and 1010 pores×cm−2, respectively. For surface modification the AAO samples were kept in a desiccator with one drop of 3ethoxydimethylsilylpropylamine at a reduced pressure of 16 mbar at room temperature for 24


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h. The samples were functionalized by depositing one bilayer of PSS and PAH inside aminosilane primed AAO. All AAO samples were coated with a thin Au film by sputtering ∼ 10 nm

gold (Edwards Sputter Coater S150B) at 1.5 - 2.0 mmHg to afford a passivation layer that prevents the polyelectrolyte from adsorbing at the surface of template,78 to increase the reflectivity and improve the RIfS signal. For kinetic measurements a custom-made flow cell for RIfS was used. For LbL PSS and PAH solutions with the same concentration of 0.5 g×L−1 in aqueous 0.1 M KCl were used. The polyelectrolyte solutions were pumped into the flow cell at a flow rate of 0.16 mL×min−1. Rinsing steps using Milli-Q water were applied in between the incubations with oppositely charged polyelectrolytes. K-phosphate buffer solution was introduced to the flow cell for 10 min before the DNA solution was injected and the surface was washed with the K-phosphate buffer again. For the SPR measurements, the SPR sensor was functionalized with a negatively charged monolayer of MHDA by immersing the sensor in 1.0 mM MHDA solution in ethanol overnight followed by rinsing with ethanol and Milli-Q water. The sensor was mounted in the flow cell of an Optrel Multiskop SPR (Optrel GbR, Sinzing, Germany). PAH (0.5 g×L−1 in aqueous 0.1 M KCl in solution) was introduced to the flow cell before washing with Milli-Q water utilizing a pump (Ismatec Laboratoriumstechnik GmbH, Wertheim-Mondfeld, Germany) with a constant flow rate 5 µL×s−1. After depositing the PAH layer K-phosphate buffer was injected into the flow cell for approx. 1 h. DNA solutions with corresponding concentrations were subsequently injected into the flow cell before the cell was washed with pure K-phosphate buffer until equilibrium was reached. The association and dissociation processes were monitored and analyzed using Optrel’s Multi.exe software, in which the angle corresponding to the minimum of the SPR reflectivity curve was measured for each stage. Binding experiments in nanopores and on the flat surface were performed in triplicate for each DNA concentration.


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For the CFM measurements, the fluorescence intensity data of DNA-ligand complexes was determined on glass cover slides (20 ×20 mm2, Menzel, VWR, Germany) at room temperature in buffered solution. The glass slides were cleaned by sonication in ethanol and water followed by UV-ozone treatment. After the functionalization with a PAH layer, spreading 5µM DNA solution on the slides for 10 min, and rinsing with K-phosphate buffer solution, the ligand solution (c = 2.5 µM) was applied to the film. After incubation for 10 min, the surface was rinsed with K-phosphate buffer solution and imaged. The ligand was excited by a pulsed laser (LDH-P-C-635B) at 635 nm. The fluorescence was detected by a single-photon avalanche diode (PD5CTC, Micro Photon Devices, Bolzano, Italy). The objective was held on a XYZ piezo controller and was scanned in XY-plane in a range of 80 × 80 µm2 with a pixel resolution of 512 × 512. The dwell time was 4.8 ms/pixel. The data were analyzed using the SymPhoTime (version 5.2.4.0, PicoQuant, Berlin, Germany) and OriginLab 9.0 software (OriginLab, Northampton, USA). The measurement was performed before and after ligand addition accompanied with images recording. The observed rate constants (kobs) were determined by fitting the adsorption kinetics obtained from SPR and RIfS experiments to the established theoretical model (eq. 2).79

∆ = ∆ ! (1 − % &'() )

(eq. 2)

Where ∆Rt is the change of the reflectivity or ∆EOT for SPR or RIfS, respectively at time t and cDNA is the DNA concentration. The measurements of the binding kinetics of the G4-DNA and 1c in liquid were performed using the confocal microscope by monitoring the fluorescence intensity of 1c upon mixing with G4-DNA. A volume of 2 µL of the dye solution (200 nM) was deposited onto a glass slide, 18 mL of the G4-DNA solution was added and mixed by pipetting the entire droplet up and down 5 times. The confocal volume was positioned above the glass surface within the


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droplet and the fluorescence intensity was recorded starting before the DNA was added. To get the apparent rate constant kobs of the reaction an exponential function was fitted to the increasing fluorescence intensity I(t) after mixing.

+(,) = +- + ∆+/1 − % &'() (0 ) 1 (eq. 3) where t and t0 are the time and the time at which the reaction started and I0 and ∆I are the initial fluorescence intensity and its change upon binding, respectively. The association rate constant (kon) and the dissociation rate constant (koff) were determined by fitting the observed rate constants according to the Langmuir model (eq. 4).80

234 = 2 5678 + 299

(eq. 4)

The maximum surface excess (Γmax) and the Langmuir constant (KL) were determined from the SPR and RIfS data using eq. 5.81 : = [B ?

;

?@ @ ]

(eq. 5)

Where Γ is the surface excess, which corresponds to the angle shift (∆θ) or the change of the effective optical thickness (∆EOT), and c is the concentration. Based on the measured value for ∆EOT the refractive index of the void pores nvoid is calculated according eq.6 and eq.7 and is used to get the volume fraction f2 of the material deposited in the pores (eq. 9) which yields the thickness l of that layer. ∆EOT is calculated as the difference of individual EOT readings and depends on L is the AAO film thickness, i.e. the length of the pores (nm), and the effective refractive index ne of the porous material ,according to equation (eq. 6): ∆EOT= 2neL

(eq. 6).

ne ih can be calculated from the Maxwell-Garnett equation (eq. 7):82 DE = D8F GH

K'LM NOK'LM I@ P I@ JH JH I@ K'LM NOK'LM I@ P JH JH

(eq. 7)


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Where: nAL2O3 is the constant refractive index of the bulk AAO (nAl2O3 = 1.70 in this work)83, nvoid is the refractive index of the material filled or deposited in the pores, and P is the porosity of the AAO pores, which can be determined based to the pore diameter and the inter pores distance calculated from the SEM images. The thickness of DNA layer in AAO substrate was calculated from the acquired RIfS spectra applying the two components Lorentz-Lorenz equation (eq. 8):84 K B

K

= QB + Q B

B

(eq. 8)

Where nv is the refractive index of the void medium inside pores (calculated from the acquired RIfS spectra applying the Maxwell-Garnet approach (eq.7),85 n1 is the refractive index of the Milli-Q water, n2 is the refractive index of the deposited material either PSS/PAH bilayer (n2 = 1.50) 86 or DNA layer (n2 = 1.46), f1 and f2 are the volume fractions of Milli-Q water and the deposited material respectively. For the SPR measurements the layer thickness l of the DNA is fitted with Fresnel equations by the Winspall software 3.2 (Max Planck Institute of Polymer Research, Mainz, Germany) depending on the reflectivity scan data using the refractive indices of different materials (Table 3).

Table 3. Refractive indices of different layers at wavelength of 633 nm. Layer

Refractive index

LSF9 Prism

1.52

Ti

2.054 a + i 3.142

Gold

0.183 b + i 3.343

MHDA

1.48c

PAH

1.56d

K-phosphate buffer

1.335e


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DNA

1.46f

a

Ref.87. b Ref. 88. c Ref. 89. d Ref. 90. e measured by refractometry. f Ref. 90. The mass deposition M of DNA on planar substrates and in AAO nanopores (ng×mm−2) was determined by eq. 9.

R =S∙U

(eq. 9)

Here, ρ is the density and l is the thickness of the deposited DNA layer. Assuming a density of DNA of 1.7 g×cm−3,91 the pore diameter after one bilayer deposition is 51 nm (as PSS/PAH bilayer thickness is 2 nm, calculated from obtained EOT value), and the pores density is 1010 pores×cm−2.

Supporting Information Fluorimetric titration of 1c with ct DNA, 1H-NMR spectra of the complex between Tel26 and ligand 1c, confocal fluorescence microscopy images of DNA-coated surfaces with 1c, SEM images of AAO, and scheme of binding kinetics.

Acknowledgements The authors gratefully acknowledge Prof. Andreas Kolb and Dipl.-Inf. Hendrik Hochstetter for inspiring discussions, Ms. Sandra Uebach for technical assistance with the NMR experiments, Professor Michael J. Sailor, University of California for kindly providing the reflectometry analysis software as well as Dr. Marc Steuber and M. Sc. Qasim Alhusaini for acquiring the SEM data. Financial support by the Deutsche Forschungsgemeinschaft (INST 221/87-1 FUGG), the European Research Council (ERC project ASMIDIAS to HS, Grant no. 279202) and the University of Siegen (incl. a Research Grant of the School and Science and Technology) is gratefully acknowledged.


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ToC graphics


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References

(1) Victorelli, S.; Passos, J. F. Telomeres and Cell Senescence – Size Matters Not. EBioMedicine 2017, 21, 14−20. (2) Kirchner, H.; Shaheen, F.; Kalscheuer, H.; Schmid, S. M.; Oster, H.; Lehnert, H. The Telomeric Complex and Metabolic Disease. Genes 2017, 8, 176. (3) Venkatesan, S.; Khaw, A. K.; Hande, M. P. Telomere Biology−Insights into an Intriguing Phenomenon. Cells 2017, 6, 15. (4). Xu, Y. Chemistry in human telomere biology: structure, function and targeting of telomere DNA/RNA. Chem. Soc. Rev. 2011, 40, 2719−2740. (5) Lu, W.; Zhang, Y.; Liu, D.; Songyang, Z.; Wan, M. Telomerase-structure, function and regulation. Exp. Cell Res. 2013, 319, 133−141. (6). Kim, N. W.; Piatyszek, M. A.; Prowse, K. R.; Harley, C. B.; West, M. D.; Ho, P. L. C.; Coviello, G. M.; Wright, W. E.; Weinrich, S. L.; Shay, J. W. Specific association of human telomerase activity with immortal cells and cancer. Science 1994, 266, 2011−2015. (7). Shay, J. W.; Wright, W. E. Telomerase therapeutics for cancer: challenges and new directions. Nat. Rev. Drug Discov. 2006, 5, 577−584. (8) Collie, G. W.; Parkinson, G. N. The application of DNA and RNA G-quadruplexes to therapeutic medicines. Chem. Soc. Rev. 2011, 40, 5867−5892. (9) Mason, P. J.; Perdigones, N. Telomere biology and translation research. Transl. Res. 2013, 162, 333−342.


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(10) Ruden, M.; Puri, N. Novel anticancer therapeutics targeting telomerase. Cancer Treat. Rev. 2013, 39, 444−456. (11) Siddiqui-Jain, A.; Hurley, L. H. DNA structure: Visualizing the quadruplex. Nat. Chem. 2013, 5, 153−155. (12) Patel, D. J.; Phan, A. T.; Kuryavyi, V. Human telomere, oncogenic promoter and 5′UTR G-quadruplex: diverse higher order DNA and RNA targets for cancer therapeutics. Nucleic Acids Res. 2007, 35, 7429−7455. (13) Phan, A. T. Human telomeric G-quadruplex: structures of DNA and RNA sequences. FEBS J. 2010, 277, 1107−1117. (14) Murat, P.; Balasubramanian, S. Existence and consequences of G-quadruplex structures in DNA. Curr. Opin. Genet. Dev. 2014, 25, 22−29. (15) Petraccone, L. Higher-order quadruplex structures. Top Curr. Chem. 2013, 330, 23−46. (16) Blackburn, E. H. Telomeres and Telomerase: The Means to the End. Angew. Chem. Int. Ed. 2010, 49, 7405−7421. (17) Greider, C. W. Telomerase Discovery: The Excitement of Putting Together Pieces of the Puzzle. Angew. Chem. Int. Ed. 2010, 49, 7422−7439. (18). Balasubramanian, S.; Hurley, L. H.; Neidle, S. Targeting G-quadruplexes in gene promoters: a novel anticancer strategy? Nat. Rev. Drug Discov. 2011, 10, 261−275.


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Fluorimetric detection of G-quadruplex DNA in solution and adsorbed

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