Study of a DNA Duplex by Nuclear Magnetic Resonance and

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A Study of a DNA Duplex by Nuclear Magnetic Resonance (NMR) and Molecular Dynamics Simulations. Validation of Pulsed Dipolar Electron Paramagnetic Resonance Distance Measurements using Triarylmethyl-Based Spin Labels Alexander Anatolyevich Lomzov, Eugeniy A. Sviridov, Andrey V. Shernuykov, Georgiy Yu Shevelev, Dmitrii V. Pyshnyi, and Elena G. Bagryanskaya J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.6b03193 • Publication Date (Web): 19 May 2016 Downloaded from http://pubs.acs.org on May 25, 2016

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The Journal of Physical Chemistry

A Study of a DNA Duplex by Nuclear Magnetic Resonance (NMR) and Molecular Dynamics Simulations. Validation of Pulsed Dipolar Electron Paramagnetic Resonance Distance Measurements using Triarylmethyl-Based Spin Labels Alexander A. Lomzov,1,2.∥ Eugeniy A. Sviridov,2,3.∥ Andrey V. Shernuykov,3 Georgiy Yu Shevelev,1,2 Dmitrii V. Pyshnyi,1,2,* and Elena G. Bagryanskaya2,3* 1

Institute of Chemical Biology and Fundamental Medicine, SB RAS, 8 Lavrentiev Avenue,

Novosibirsk 630090, Russia 2 3

Novosibirsk State University, Novosibirsk 630090, Russia N.N. Vorozhtsov Novosibirsk Institute of Organic Chemistry, SB RAS, 9 Lavrentiev Avenue,

Novosibirsk 630090, Russia Email: [email protected]

ACS Paragon Plus Environment


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ABSTRACT Pulse dipole−dipole electron paramagnetic resonance (EPR) spectroscopy (double electron−electron resonance [DEER] or pulse electron–electron double resonance [PELDOR] and double quantum coherence [DQC]) allows for measurement of distances in biomolecules and can be used at low temperatures in a frozen solution. Recently, the possibility of distance measurement in a nucleic acid at a physiological temperature using pulse EPR was demonstrated. In these experiments, triarylmethyl (TAM) radicals with long memory time of the electron spin served as a spin label. In addition, the duplex was immobilized on modified silica gel particles (nucleosilDMA); this approach enables measurement of interspin distances close to 4.5 nm. Nevertheless, the possible influence of TAM on the structure of a biopolymer under study and validity of the data obtained by DQC are debated. In this paper, a combination of molecular dynamics (MD) and nuclear magnetic resonance (NMR) methods was used for verification of interspin distances measured by the Q-band DQC method. NMR is widely used for structural analysis of biomolecules under natural conditions (room temperature and an aqueous solution). The ultraviolet (UV) melting method and thermal series 1H NMR in the range 5–95 °C revealed the presence of only the DNA duplex in solution at oligonucleotide concentrations 1 µM to 1.1 mM at temperatures below 40 °C. The duplex structures and conformation flexibility of native and TAM-labeled DNA complexes obtained by MD simulation were the same as the structure obtained by NMR refinement. Thus, we showed that distance measurements at physiological temperatures by the Q-band DQC method allow researchers to obtain valid structural information on an unperturbed DNA duplex using terminal TAM spin labels.


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INTRODUCTION The development of approaches to structural analysis of biologically important molecules, in particular, of keystone biopolymers, proteins, and nucleic acids and their complexes is important for understanding of their functions. In the cases when crystal structure of biomolecules cannot be determined (and therefore X-rays analysis is not applicable), magnetic resonance methods (nuclear magnetic resonance [NMR] and electron paramagnetic resonance [EPR]) as well as fluorescence resonance energy transfer (FRET) are useful.16 NMR spectroscopy enables experiments in an aqueous buffered solution and studies on objects under conditions mimicking natural biological systems as well as exploration of the dynamics of their interaction with other molecules. This knowledge is important for elucidation of their functions. Multidimensional NMR is used to overcome experimental limitations for torsional angles and distances between magnetically active nuclei up to 0.6 nm. The latter can lead to an ambiguous readout from biological molecules with tertiary structure. In such cases, the distances between parts of biomolecules located at long distances can be determined using other methods, in particular, pulse electron–electron double resonance (PELDOR)7 or double electron−electron resonance (DEER)8 and double quantum coherence (DQC)9 spectroscopy. To apply this approach, investigators use biopolymers that possess spin labels (usually, nitroxide radicals) at desired locations. EPR with single spin labeling of biomolecules provides structural and dynamic information on the biopolymer fragment containing the label, in particular, on accessibility of the label to external paramagnetic probes from the solvent, on polarity of the molecular environment, and mobility of the label (for a review, see 3,4,5,6,1013). PELDOR/DQC spectroscopy is widely applied to systems with two spin labels introduced at desired locations of one biopolymer or different biopolymers forming a specific complex. PELDOR/DQC allows for measurement of intramolecular and intermolecular interspin distances when their values fall into the range between 2 and 8 nm14,1617 and has been fruitfully used for research on proteins and nucleic acids as well as their complexes.234,18 An important advantage of PELDOR/DQC is higher (1000-fold) sensitivity in comparison with NMR and applicability to heterogeneous and nontransparent samples. The combination of NMR and PELDOR/DEER yields the best results and allows researchers to solve complicated biological problems.1 Until recently, PELDOR/DQC required freezing the sample and performing the experiments at liquid-helium or liquid-nitrogen temperatures because high mobility of spin labels leads to averaging of the anisotropic dipole–dipole interaction to zero, and as a result, to disappearance of oscillation in



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the decay of electron spin echo time dependence. This state of affairs leads to general uncertainty in matching of natural- and frozen-state structures. Moreover, freezing of aqueous solutions generally requires addition of cryoprotectants (such as glycerol), which may also affect the distance distributions and conformational equilibria. Recently, various approaches enabling distance measurements at room and physiologically relevant temperatures were developed. These approaches are based on the use of spin labels with long relaxation time of the electron spin and on immobilization of a biomolecule by coupling with Sepharose,25 electrostatic interaction with modified silica gel,26 or embedding in a trehalose matrix27,28. For the most common nitroxide spin labels, such as 1-oxyl-2,2,5,5- tetramethyl-3pyrroline-3-(methyl)methanethiosulfonate (MTSL), the value of phase memory time is too short to perform PELDOR/DQC experiments at temperatures higher than 80 K because of fast rotation of the methyl groups at rates comparable to anisotropy of hyperfine couplings on methyl protons.29 The use of triarylmethyl (TAM) radicals30,31 or spirocyclohexane-substituted nitroxides28,32,33 with longer memory time of the electron spin34,35 allows for measurement of distances at liquid nitrogen and even physiologically relevant temperatures.25,26 The TAM radical has much longer relaxation time in comparison with nitroxides34 and is a promising alternative to nitroxide spin labels for dipolar EPR spectroscopy. Recently, the distances as long as 4.6 nm were measured at 37 °C by pulse EPR using a DNA duplex doubly spin-labeled with TAM and immobilized on the ion exchange sorbent NucleosilDMA.26 A disadvantage of TAM radicals when used as spin labels is a substantially larger size in comparison with nitroxides as well as hydrophobicity. For these reasons, the possible influence of TAM on the structure of a biopolymer under study as well as validity of the information obtained by pulse EPR are currently debated. In this paper, a combination of molecular dynamics (MD) and NMR methods was used for verification of interspin distances measured by the Q-band DQC method. A native DNA duplex and a doubly triarylmethyl-labeled analog were used as model objects. To confirm formation of only the DNA duplex by the oligonucleotides, we utilized the thermal denaturation method and the imino proton NMR analysis.

MATERIALS AND METHODS Oligonucleotide Synthesis, Purification, and Sample Preparation. Oligonucleotides D1 (5′CACGCCGCTG-3′) and D2 (5′-CAGCGGCGTG-3′) were synthesized in accordance with standard


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protocols of phosphoramidite chemistry from commercially available monomers (Glen Research, USA) and were purified by ion exchange high-performance liquid chromatography (HPLC) [Polysil 300-CA (10 × 250 mm, 17 µm; ICBFM SB RAS, Russia), 0.0–0.3 M KH2PO4 in 30% aqueous acetonitrile], and followed by reverse-phase HPLC [ZORBAX Eclipse XDB-C18 (4.6 × 150 mm, 3.5 µm; Agilent Technologies, USA), 0–50% aqueous acetonitrile containing 0.02 M tetraethylammonium acetate]. Concentration of the oligonucleotides was calculated from their ultraviolet (UV) spectrum absorbance (Shimadzu UV-VIS 2100, Japan) by means of molar extinction coefficients at 260 nm published elsewhere.36 The oligonucleotides were also subjected to successive passage through a small column packed with Dowex 50 (Na+) and Chelex 100 (Na+) resins for NMR experiments. Then, the samples were lyophilized twice to dryness from D2O and dissolved in 0.6 ml of a buffer (100 mM NaCl, 10 mM NaH2PO4, 200 µM EDTA sodium salt, pH 7.2) in D2O (99.9999% D, Sigma-Aldrich, USA) or the mixture H2O/D2O (9:1, v/v). Samples contained the DNA duplex at 1.1 mM. Thermal Denaturation of DNA Duplexes. Thermal stability of DNA duplexes was studied by the UV melting method. Thermal denaturation or renaturation experiments were carried out on a Cary 300 Bio spectrophotometer equipped with a six-cell Peltier thermostated cuvette holder (Varian, Australia). Melting curves were recorded at 260 and 270 nm within the temperature range 5–95 °C in an aqueous buffer consisting of 100 mM NaCl, 10 mM NaH2PO4, and 200 µM EDTA sodium salt (pH 7.2). The rate of temperature change was 0.5 °C/min. Duplexes were formed by stoichiometric mixing of the D1 and D2 oligonucleotides. Thermodynamic parameters of the non–self-complementary duplex formation were determined by fitting of the melting curves or by the concentration method in accordance with two-state models described previously.37 In the second case, equation (1) was used to determine the thermodynamic parameters:

(1), where Tm is melting temperature, ∆H0 and ∆S0 are hybridization enthalpy and entropy, respectively, CT is the total oligonucleotide concentration, and R is a gas constant.



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NMR Experiments. The NMR spectra were recorded on a Bruker AV-600 spectrometer (1H: 600.30 MHz;

31

P: 243.1 MHz) at the Multi-Access Chemical Service Center, SB RAS. The data were

processed using TopSpin 3.2 (Bruker, Germany), and the cross-peaks were assigned using CcpNmrAnalysis 2.3.1.38 Proton chemical shifts were referenced to an internal standard, 4,4-dimethyl4-silapentane-1-sulfonic acid (DSS) at 0.00 ppm; 31P chemical shifts were indirectly referenced to the 1

H standard.39 Spectra of nuclear Overhauser effect spectroscopy (NOESY; mixing time 100, 200, and

400 ms) and total correlation spectroscopy (TOCSY; mixing time 60 ms) were recorded in D2O at 33.3 and 15.3 °C, respectively. For the NOESY experiments, datasets of 4096 (t2) × 512 (t1) complex points were acquired with 16 scans per t1 increment and the spectral width of 6313.13 Hz. For the TOCSY experiment, datasets of 4096 (t2) × 512 (t1) complex points were acquired with 8 scans per t1 increment and the spectral width of 6355.93 Hz. Watergate-NOESY experiments40 (mixing time 100, 200, and 400 ms) in the mixture H2O:D2O (9:1) were conducted at 15.3 °C. Datasets of 4096 (t2) × 512 (t1) complex points were acquired with spectral width of 15000.0 Hz in both dimensions and 16 scans per t1 increment. Assignment of Proton Chemical Shifts. Assignment of the majority of 1H chemical shifts was based on the set of NOESY experiments (mixing time 100, 200, and 400 ms) and TOCSY experiments (mixing time 60 ms). Sequential assignment of nonexchangeable protons was based on the throughspace connectivity with H1′ and H2′/H2′′ protons in the 1H-1H NOESY spectra.41 All expected H6/H8base and H1′/H2′/H2′′-ribose connectivity values were identified in the NOESY spectra. Experimental values of assigned chemical shifts are summarized in the table S1 (Supporting Information). MD Simulation of TAM-Labeled and Native DNA Duplexes. The MD simulations were performed in the Amber 12 software.42 Library files of the TAM spin label with a linker and an internal cytosine nucleotide connected to TAM were obtained earlier.26 Fifteen initial structures were obtained by the cluster analysis of a 210-ns MD trajectory of the TAM-labeled duplex. The structure of triarylmethyllabeled D1-D2 duplex and initial conformations showed in Figure S6 (Supporting Information). The initial structures of the native duplex were prepared from TAM-labeled complexes by removing the terminal spin labels with the linker. MD simulations were conducted using the ff99bsc0 force field in an explicit solvent. A TIP3P water model with a radius of 12 Å was applied. To neutralize the net charge, 18 Na+ ions were added. Periodic boundary conditions, an isothermal–isobaric ensemble with the constant number of particles (NTP) with isotropic position scaling at 300 K, and the Andersen


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temperature coupling scheme were used. The particle mesh Ewald (PME) method with a cutoff of 8 Å and the SHAKE algorithm for bonds involving hydrogen were also employed. The time step was 2 fs, and the translational and rotational motion of the mass center were removed every 1 ps. The total 1.5µs trajectory length for native or modified complexes was obtained (100 ns for every initial structure). NMR volume and distance restraints were used for subsequent refinement of the structure. Peak volumes in two-dimensional (2D) NOESY spectra (100, 200, and 400 ms, 15.3 °C) were integrated by the “box sum” method in CcpNMR Analysis.43 Their values served as flat-bottom restraints with bounds of ±10% of the volume, and the overall scaling factor was set to 4.37 × 10−5; molecule rotational tumbling time was 3 ns, and the correlation time for methyl groups was 40 ps. Сross-peak volumes were also converted into interproton distances in the IRMA2 software.44 We utilized scaling with reference peaks, and H5-H6 cytosine cross-peaks served as a reference with the distance of 2.45 Å. Overall correlation time was set to 3.50 × 10−9 s, and the additional leakage rate was 0.8 s−1 for IRMA2 calculations. The lower and upper bounds of the restraints were set to ±15% of the calculated NOE distance. For those distances that IRMA2 failed to calculate, we set lower and upper bounds to 1.8 and 7.0 Å, respectively. Watson–Crick hydrogen bonding of all base pairs (except terminal ones) was identified by NOE correlations G(N1–H)–C(NH2) or A(H2)–T(N3–H) and significant chemical shifts of imino protons. H-bond restrains that are based on the standard B-form DNA geometry were plugged into calculations, with the tolerance of ±0.1 Å. The IRMA2 software was used to verify the resulting structures by back-prediction of NOESY spectra and by comparing the empirical and back-predicted spectra. The value of r6-weighted R-factor (IRMA2 R5 factor) was selected as a criterion of structure consistency. Structural calculations were performed using the SANDER module of Amber 12. The generalized Born implicit solvent model with the equivalent of 0.1 M 1-1 ions and the Andersen temperature coupling scheme were used. The PME method with a cutoff of 8 Å was applied. The B-form D1–D2 duplex was heated up beforehand to 1000 K for 0.5 ns with the time step of 0.0005 ps and was equilibrated for 0.5 ns. Watson–Crick H-bond distance restraints were applied (force constant 1 kcal/[mol⋅Å2]). The last 400 ps of the trajectory were used to obtain 10 structures with the interval of 40 ps. These were initial structures for MD simulations with restraints (Supporting Information Figure S5). An increasing force constant from 1 to 35 kcal/mol for distance and volume restraints was



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maintained for the subsequent duplex cooling to 300 K for 1 ns. The productive MD trajectory of DNA was simulated with the force constant of 35 kcal/mol for distance and volume restraints. Trajectory analysis was performed using the cpptraj tool of Amber 12. Hierarchical cluster analysis was conducted for productive MD trajectory analysis of the DNA duplexes without terminal base pairs, and a random sieve of 100 was applied. Molecular graphics were prepared by means of the UCSF Chimera package.45 RESULTS Thermal Denaturation of the DNA Duplex. To ensure that oligonucleotides D1 and D2 formed only the DNA duplex in solution, thermal denaturation of the DNA duplex at various concentrations of the complex was performed (Supporting Information Figure S1). Denaturation curves were reversible (Supporting Information Figure S2). The enthalpy and Gibbs free energy changes that were obtained at 260 and 270 nm by fitting of the melting curves using the two-state model did not differ by more than 8% and 3%, respectively. Thermodynamic parameters at different concentrations obtained by the fitting method were identical and close to those calculated by the concentration method (Table 1). A linear dependence of 1/Tm on Ln(CT/4) was observed (R2 = 0.998, Supporting Information Figure S3). In addition, thermal denaturation of the single-stranded oligonucleotides was carried out. We did not observe any transitions during heating of the single-stranded oligonucleotides (Supporting Information Figure S2). Taken together, these data confirm formation of the duplex according to the two-state model, in other words, a system consisting of only one species of the double-stranded (ds) bimolecular complex or single-stranded oligonucleotides at low and high temperatures, respectively.46 Table 1. Thermodynamic parameters of DNA duplex formation at different concentrations of the D1– D2 complex obtained by fitting of UV melting curves, by the concentration method, and by fitting the temperature dependence of the chemical shift of proton H7 of nucleotides T19 and T9. C, µM

2 5 10 20

∆H0,

∆S0,

∆G037,

kcal/mol

cal/[mol⋅K]

kcal/mol

-63.7 ± 1.7 -71.8 ± 6.0 -76.7 ± 6.2 -70.5 ± 4.4

-167.4 ± 5.5 -191.8 ± 18.0 -207.3 ± 19.1 -188.3 ± 13.3

-11.8 ± 0.1 -12.3 ± 0.4 -12.4 ± 0.3 -12.1 ± 0.3


Tm, °C

51.7 ± 0.6 54.8 ± 0.3 56.2 ± 0.3 58.5 ± 0.2

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50 -72.0 ± 0.8 -192.6 ± 2.1 -12.2 ± 0.1 61.4 ± 0.2 100 -72.1 ± 1.0 -193.1 ± 2.9 -12.2 ± 0.1 63.5 ± 0.2 200 -74.5 ± 0.8 -200.4 ± 2.4 -12.3 ± 0.1 65.3 ± 0.2 2200* -73.6± 3.0 -197.8 ± 10.4 -12.3± 0.1 73.8± 0.3 # 2200 -71.6 ± 4.0 -191.6 ± 12.4 -12.2 ± 0.2 74.2 ± 0.3 2200& -64.1 ± 1.3 -169.2 ± 3.9 -12.3 ± 0.1 75.0 ± 0.3 The data presented as mean ± standard deviation (SD). *Obtained by the concentration method in accordance with equation (1); #the thermodynamic parameters were calculated as the average of the values obtained by fitting and concentration methods; &obtained by fitting of the NMR data. Analysis of DNA Duplex Structure by NMR Spectroscopy. The melting temperature of the DNA duplex calculated using UV melting data and equation (1) at concentration of NMR experiments should be ~74 °C (Table 1). To confirm the duplex formation, a temperature series of 1H NMR imino proton

spectra

was

recorded.

Figure 1. An 1H NMR spectrum of the imino proton region of the DNA duplex (a). Temperature dependence of the imino proton region of the 1H NMR spectra (b). The data in Figure 1 show two main regions of imino protons: thymine imino protons in the range 14.1–14.2 ppm and guanine in the range 12.6–13.2 ppm. The imino proton peaks corresponding to the terminal guanine residues were not observed due to the fast exchange with the solvent because of frying.47 Analysis of the temperature series of the imino proton region from 1H NMR spectra showed a decrease in signal magnitudes that correspond to disruption of the Watson–Crick base pairs and to duplex denaturation. A rise in temperature from 6 to 65 °C revealed comparable stability of allGCbase pairs. In contrast, the signal of preterminal thymidines (T9 and T19) decreased faster with the



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temperature increase because of the intensifying proton exchange with the solvent. The changes in the guanine’s part of the spectra correspond to the whole-complex stability. To confirm formation of only the DNA duplex under NMR conditions, a 1H NMR temperature series was recorded. Analysis of the chemical shift of the H7 proton of T19 and T9 (dependence on temperature) showed a shape transition (Figure 2, Supporting Information Figure S4 ). A comparison of the denaturation curve calculated using the thermodynamic parameters obtained by the UV melting method and by means of the NMR data is shown in Figure 2. The two types of data are in excellent agreement. A quantitative comparison of the thermodynamic parameters calculated by fitting NMR data and by optical measurements correlates very well (Table 1). This finding confirms the presence of only the DNA duplex at 37 °C in the concentration range from 1 µM to 1.1 mM and enables comparison of the data obtained by the NMR and EPR methods.

Figure 2. Temperature dependence of the chemical shift of the H7 protons of T19 (red circles) and T9 (blue diamonds) derived from 1H NMR spectra and calculated using average thermodynamic parameters (Table 1) obtained by the UV melting method (lines) at duplex concentration 1.1 mM. Resonance Assignment of Nonexchangeable Protons. Assignment of the majority of 1H chemical shifts was based on the set of NOESY experiments (mixing times 100, 200, and 400 ms) and TOCSY experiments (60-ms mixing time). Signals H6-H2′ and H6-H2′′ of the first cytosines of both strands were easily identified by their specific chemical shifts and by the absence of NOE with the next residues. Sequential assignment of nonexchangeable protons was based on the values of through-space intra- and internucleotide connectivity with H1′ and H2′/H2′′ protons in the 1H-1H NOESY spectra (Figure 3). The use of two pathways (via H1′ and H2′/H2′′ protons) allowed us to avoid ambiguities in


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proton assignment. Stereospecific assignment of the H2′/H2′′ protons was attained according to the rule that for H1′–H2′′, NOE is stronger than for H1′–H2′. All expected connectivity values of the H6/H8 nucleobase and H1′/H2′/H2′′ ribose were identified in the NOESY spectra. Analysis of intraresidue and sequential NOEs between H1′, H2′, H2′′, H3′ and base H6/H8 revealed the B conformation of the DNA double helix.

Figure 3. Sequential assignment of nonexchangeable protons in 1H-1H NOESY spectra (mixing time 400 ms) obtained via H1′ (top) and H2′/H2′′ protons (bottom) for D1 (green solid line) and D2 strands (red dashed line).



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In the NOESY spectrum acquired in the H2O/D2O mixture (9:1), we observed hydrogen-bonded protons from all G, T, and C residues except for the terminal ones. This result indicates the presence of the Watson–Crick bonding for the corresponding base pairs. The absence of signals of the terminal nucleobases caused by frying.47 Imino protons of residues T9 and T19 were assigned on the basis of the correlation between imino protons of thymines and H2 protons of A12 and A2, respectively. The chemical shifts of imino protons and the corresponding NOE connectivity pattern were in agreement with the canonical Watson–Crick base pairing, and all our NMR data were consistent with unperturbed B-DNA structure. The peak volumes and calculated distances are summarized in Table S2 (Supporting Information). Refinement of the D1/D2 DNA Duplex Structure. Different initial structures were used for MD simulations, with distance restraints obtained from the NMR data. After cooling and equilibration of the DNA duplexes with the NMR restrains, 10 series of 100-ns trajectories were obtained and used in the subsequent analysis. The cluster analysis of the whole MD trajectories allows us to distinguish five conformations of the DNA duplex. Two of them were the most prevalent during the simulation and occupied 70% and 20% of the trajectory. All structures matched the distance restraints well, none of them showed an intraproton distance difference of more than 0.5 Å from restraint bonds, whereas the values of r6-weighted R-factor were no more than 0.0032 for all peaks. Back-calculated volumes of NOESY cross-peaks also correlated well with empirical cross-peak volumes. The structural statistics and cluster analysis data are summarized in Table 2 and Table S3 (Supporting Information), respectivelly. Table 2. Structural parameters of five conformations of the DNA duplex. r6-weighted RSimulation Energy penalty Pearson R factor* RMSD# factor # population, Volume, Distance, Intra Inter Bond Bond All Inter% All kcal/mol kcal/mol residual residual length, Å angle, Deg peaks residual 1 70.1 305.2 96.1 0.876 0.91 0.734 0.0203 3.47 0.0027 0.0048 2 19.8 336.3 162.7 0.842 0.869 0.674 0.02 3.73 0.0029 0.005 3 9.7 360.5 171 0.833 0.836 0.747 0.0132 2.98 0.0029 0.005 4 0.2 340.4 164.4 0.859 0.901 0.562 0.0208 3.89 0.0032 0.0056 5 0.3 287.7 102.6 0.877 0.947 0.621 0.0198 3.41 0.0028 0.0056


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*

Pearson R factor calculate for peak volumes obtained in NMR experiments and calculated in MD simulation; # RMSD values calculated from ideal geometry.

To summarize all the NMR results, it was shown that the duplexes are B-form DNA with canonical Watson–Crick base pairs. MD Simulations of Native and TAM-labeled DNA Duplexes. To verify the interspin distances measured by the Q-band DEER method, the 1.5- µs MD simulations of the native and TAM-labeled DNA duplexes were performed. The distance distributions presented in Figure 4a show good agreement. The average interspin distances obtained by single Gaussian fitting were 45.7 ± 1.6 and 45.4 ± 2.2 Å, respectively (mean ± SD). The slight slide of the maximum of the MD distribution into the long-distance range may be caused by inaccuracy of the simulation, in particular, by the non-ideal force field. To test this hypothesis, the unmodified dsDNA was simulated in the same manner as the TAM-labeled duplex was. Then, its structure was compared with the data obtained by NMR spectroscopy. During analysis of the influence of the spin labels on duplex structure, the distances between the most faraway C5′-atoms of the 5′-terminal nucleotides of oligonucleotides D1 and D2 were determined. The distance distributions are presented in Figure 4b. The maxima and widths of the distance distributions were very close. The mean distance values and widths of distributions determined along 1.5-µs trajectories were 31.4 ± 2.1 Å for native and 31.2 ± 2.2 Å for modified duplexes. This result means that there is no significant effect of the TAM labels on the C5′–C5′ distance and thus on the whole duplex structure and duplex dynamics.



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Figure 4. (a) The distribution of interspin distance probability in the TAM-labeled DNA duplex obtained by analysis of a 1.5-µs trajectory (thick curve), MD trajectory, and by the DQC method (thin curve, adapted from 26). (b) The C5′–C5′ distance of the 5′-terminal nucleotides obtained from the MD simulation of the TAM-labeled DNA duplex (red), native DNA duplex (blue, dashed), and native DNA duplex with NMR restraints (green, dash-dotted). A comparison of C5′–C5′ distance distributions obtained for the native D1–D2 duplex obtained with and without NMR restraints is shown in Figure 4b. The maxima and width of the distribution obtained in MD simulations with restraints is 31.4 ± 2.0 Å, which is close to the results of unrestrained simulations. To compare TAM-labeled, native, and NMR-refined structures, a cluster analysis of each MD trajectory was performed. The average distance between structures in the most populated clusters was 1.5 Å in all cases (Supporting Information Figure S7 and Table S3). A comparison of the most representative structures in each type of MD trajectory is shown in Figure 5. The root-mean square deviation (RMSD) between heavy atoms of the duplexes with excluded terminal base pairs was 0.7 Å. Together with the C5′–C5′ distance distributions, these data mean that the structure and conformational flexibility of native and spin-labeled D1–D2 duplexes obtained by the MD simulations with and without NMR restraints are the same.


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Figure 5. Comparison of structure of TAM-labeled (red), native (blue), and native NMR-restrained D1/D2 DNA duplexes (green) obtained by MD simulations.

DISCUSSION The temperature dependence of NMR spectra of the native DNA duplex can be compared with thermal denaturation data obtained by optical melting. Because of the high concentration of the nucleic acids (1.1 mM) in the NMR experiments, we constructed a thermal denaturation curve from the average thermodynamic parameters obtained in the UV-melting experiments (Table 1). A comparison of the thermal denaturation data with the registered optical density and the imino proton chemical shift of 1H NMR spectra clearly shows the presence of only the perfectly matched DNA duplex formed by oligonucleotides D1 and D2 in solution (Figure 2). Good agreement between the two types of data confirms that the denaturation of the nucleic acid complex conforms to the two-state model.46 This means that at temperatures below 40 °C, sample consists practically only the matched DNA duplex is present in the system at oligonucleotide concentrations 1 µM to 1.1 mM (Supporting Information Figure S1). The 1.5-µs trajectory length is more ergodic than the previously published one26 obtained using 210-ns trajectory length. Therefore, these data are more suitable for comparison with the empirical results



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obtained by NMR and Q-band DEER methods. The most remote C5′–C5′ atoms of the 5′ nucleotides of D1 and D2 were selected to compare conformational flexibility of the native and spin-labeled duplexes by MD simulation. For TAM-labeled, unrestrained native, and NMR-restrained native duplexes, the width of the distance distribution was 2.3, 2.4, and 2.0 Å, respectively. These values are close to the width of interspin distance distribution produced by the MD simulation (1.6 Å) and by pulsed EPR spectroscopy (2.2 Å). The slight broadening of the EPR distribution may be caused by the spin delocalization in the TAM labels48 or by inaccuracy of the MD simulation. The MD simulations results obtained using parmbsc0 force field in microsecond time range indicate some deviations from experimental data.495051 At the same time, the width of the interspin distance distribution obtained from Q-band DQC measurements has some uncertainty, as is usual for solutions of ill-posed problems, and therefore the observed deviations in the distribution shapes are quite typical.52 Analysis of positions of the spin labels during the simulation shows preferentially tightly interacting spin labels with terminal base pairs. Altogether, these results mean that the width of the EPR distribution (in the case of the TAM labels connected to the DNA duplex via a piperazine linker to the terminal nucleotides) is limited by duplex flexibility. For comparison of nucleic acid structures of D1–D2 and TAM-D1–TAM-D2 complexes, we selected the central part without terminal base pairs and spin labels. A cluster analysis of all three types of simulations shows that in each trajectory, only one structure is dominant. Its proportion in the case of TAM-labeled duplexes, native duplexes, and duplexes obtained with NMR refinement was 84%, 96%, and 70%, respectively. Comparison of the most probable conformations of the TAM-labeled structure, native structure, and structure with NMR restraints (Figure 5) and RMSD values for heavy atoms less than 1 Å all confirm similarity of the DNA duplex structures of the native D1–D2 complex and TAMlabeled D1–D2 complex. Moreover, the average RMSD value for all 15 duplexes obtained by cluster analysis in three simulation series was 2.0 Å (see the RMSD map in Supporting Information Figure S7). Thus, the structures of the D1–D2 DNA duplexes were the same throughout the MD simulations. This finding confirms the absence of the influence of TAM labels on duplex structure that was detected previously by the circular dichroism method.26 Our data clearly show that using TAM labels introduced at 5′ ends of the D1 and D2 oligonucleotides does not perturb the structure of the DNA duplex and could be used for distance measurement by the Q-band DEER method. CONCLUSION


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In this work, we compared the structure of native and TAM-labeled DNA duplexes by means of NMR analysis and MD simulations. To confirm that the system contains only one species and conformation of the DNA complex, we performed thermal denaturation at various duplex concentrations by the UV melting method and 1H NMR. Both approaches yielded the same results on thermal stability of native dsDNA and revealed existence of only one type of complex in solution at various oligonucleotide concentrations. The MD simulation of the native DNA duplex without restraints produces the same results as in the case of NMR restraints. Structural analysis of TAM-labeled complexes obtained by computer simulations revealed that triarylmethyl-based spin labels does not perturb either the structure of the DNA duplex or the preferential position of the spin label in the tight interaction with a terminal base pair. Moreover, the structural flexibility of the modified and native complexes simulated with and without NMR restraints is identical. In addition, the close values of the interspin distances determined by the Q-band DQC method and calculated with MD confirm reliability of measurement of the spinspin distance. In summary, the previously proposed protocol for distance measurement at physiological temperatures by the Q-band DEER method allows us to obtain correct structural information on an unperturbed DNA duplex. ASSOCIATED CONTENT Supporting Information The details of the thermal denaturation studies, 1H NMR, NMR restraints, initial structures for MD simulation, and DNA structure comparison. AUTHOR INFORMATION Corresponding Author * E-mail: [email protected], [email protected] Author Contributions ∥Alexander

A. Lomzov and Eugeniy A. Sviridov contributed equally.

Notes The authors declare that they have no competing financial interests.



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ACKNOWLEDGMENTS This work was supported by Russian Science Foundation (No. 14-14-00922), in particular NMR measurements and MD simulations . D.V.P thanks RFBR (No.16-04-01029) and MES RF (agreement no. 14.B25.31.0028.) for support of DNA synthesis and thermodynamic studies of DNA duplex by UV-melting technique, respectively. REFERENCES

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Figure 1. An 1H NMR spectrum of the imino proton region of the DNA duplex (a). Temperature dependence of the imino proton region of the 1H NMR spectra (b). 218x88mm (300 x 300 DPI)


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Figure 2. Temperature dependence of the chemical shift of the H7 protons of T19 (red circles) and T9 (blue diamonds) derived from 1H NMR spectra and calculated using average thermodynamic parameters (Table 1) obtained by the UV melting method (lines) at duplex concentration 1.1 mM. 82x61mm (300 x 300 DPI)



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Figure 3. Sequential assignment of nonexchangeable protons in 1H-1H NOESY spectra (mixing time 400 ms) obtained via H1′ (top) and H2′/H2′′ protons (bottom) for D1 (green solid line) and D2 strands (red dashed line). 82x147mm (300 x 300 DPI)


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Figure 4. (a) The distribution of interspin distance probability in the TAM-labeled DNA duplex obtained by analysis of a 1.5-µs trajectory (thick curve), MD trajectory, and by the DQC method (thin curve, adapted from 26). (b) The C5′–C5′ distance of the 5′-terminal nucleotides obtained from the MD simulation of the TAM-labeled DNA duplex (red), native DNA duplex (blue, dashed), and native DNA duplex with NMR restraints (green, dash-dotted). 149x71mm (300 x 300 DPI)



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Figure 5. Comparison of structure of TAM-labeled (red), native (blue), and native NMR-restrained D1/D2 DNA duplexes (green) obtained by MD simulations. 157x143mm (300 x 300 DPI)


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Table of Content 157x143mm (300 x 300 DPI)


Study of a DNA Duplex by Nuclear Magnetic Resonance and

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