XSL•FO - HPER Department

NAR29_14.book(gke425.fm)© 2001 Oxford University Press Nucleic Acids Research, 2001, Vol. 29, No. 14 2973–2985
Solution structure of a DNA duplex with a chiral alkyl phosphonate moiety R. Soliva, V. Monaco1, I. Gómez-Pinto2, N. J. Meeuwenoord1, G. A. Van der Marel1, J. H. Van Boom1, C. González2 and M. Orozco*
Departament de Bioquímica, Universitat de Barcelona, C/. Martí i Franquès 1-11, 08028 Barcelona, Spain, 1Gorlaeus Laboratories, Leiden University, Leiden, The Netherlands and 2Instituto de Estructura de la Materia, CSIC, C/. Serrano 119, 28006 Madrid, Spain
Received as resubmission April 26, 2001; Revised and Accepted May 21, 2001 PDB nos 1iek, 1iey
ABSTRACT
The solution structures of two DNA decamers of sequence d(CCACCpxGGAAC)·(GTTCCGGTGG) with a chiral alkyl phosphonate moiety (px) have been determined using NMR and restrained molecular dynamics simulations and compared with the solution structure of the unmodified duplex. The 1H NMR spectra of two samples with pure stereochemistry in the modified phosphate have been assigned. The structures of both diastereoisomers, as well as the unmodified control duplex, have been determined from NMR-derived distance and torsion angle constraints. Accurate distance constraints were obtained from a complete relaxation matrix analysis of the NOE intensities. The structures have been refined with state of the art molecular dynamics methods, including explicit solvent and applying the particle mesh Ewald method to properly evaluate the long-range electrostatic interactions. In both cases, the calculations converge to well-defined structures, with RMSDs of ∼1 Å. The resulting structures belong to the general B family of DNA structures, even though the presence of the alkyl phosphonate moiety induces some slight displacement to the A-form in the neighborhood of the modified phosphate. Partial neutralization of this phosphate and the steric effect of the alkyl moiety provoke moderate bending in the DNA. This effect is more pronounced in the S diastereoisomer, where the alkyl group points inwards to the double helix.
INTRODUCTION
The phosphate group is the polymerizing unit of natural nucleic acids, joining the O5′ and O3′ groups of ribose and 2′-deoxyribose. Modifications of the phosphate group induce important changes in the structure and stability of nucleic acids
(1,2). Thus, analysis of more than 100 DNA–RNA hybrids where one phosphate group was substituted by different chemical groups (1) shows that, in most cases, the resulting hybrids were less stable than the parent oligonucleotide. However, in a few cases modified nucleic acids containing non-phosphate linkages can be very stable (1–3). Clear examples are DNA–RNA hybrids containing 2′-substituted -CH2-CO-NH-CH2- linkages or methylene(methylimino) derivaties (1). Stable duplexes are also obtained when the phosphate group is replaced by, among others (1,2), phosphorothioate (4) or phosporamidate (5). Other well-studied examples of very stable nucleic acid structures are peptide– nucleic acids (PNA), where the phosphate linkage is replaced by non-natural peptide groups (6–9).
Among the different groups that can substitute the phosphate (p) linkage, alkyl phosphonates (px) are of especial interest (10–13). The substitution of phosphate by alkyl phosphonate leads to a neutral linkage, which is expected to introduce important alterations into the nucleic acid structure (2,12–15). Thus, electrophoresis experiments by Maher and co-workers (12,13) with DNA containing px derivatives placed in-phase along a DNA duplex suggest that px linking units induce curvature in the DNA. The authors raised the hypothesis that this px-induced curvature can in fact mimic that occurring in vivo when histones of the nucleosomes neutralize the charges of a few DNA phosphates (13).
In addition to DNA packing, a detailed representation of neutralization-induced bending may be crucial for an understanding of protein-induced bending of the DNA double helix (16–20). Furthermore, knowledge of the basis of chemically induced bending is crucial for an understanding of the signals triggering the action of DNA repair systems (21–23), since large localized bending is known to be recognized and repaired by DNA excision repair mechanisms (21).
In this paper we present the solution structure of a duplex DNA with a p→px substitution determined by NMR and state of the art restrained molecular dynamics simulations. Analysis of the structures and comparison with the unmodified duplex provides useful information on the basis of the chemical- and neutralization-driven bending of DNA.
*To whom correspondence should be addressed. Tel: +34 93 4021719; Fax: +34 93 421219; Email: [email protected] Correspondence may also be addressed to C. Gonzalez. Tel: +34 91 5619400; Fax: +34 91 5642431; Email: [email protected]
2974 Nucleic Acids Research, 2001, Vol. 29, No. 14
MATERIALS AND METHODS
Synthesis and purification of the oligonucleotides
Solid phase synthesis of the modified oligonucleotide 5′-d(CCACC*GGAAC)-3′ and of its complementary strand 5′-d(GTTCCGGTGG)-3′ was performed on a Pharmacia LKB Gene Assembler using the phosphoramidite method. Unmo- dified, protected 2′-deoxyribonucleoside phosphoramidites were purchased from PerSpective Biosystem and used as 0.1 M solutions in dry acetonitrile (DNA synthesis grade on 4 Å molecular sieves; Biosolve). The modified base 5′-O-(4,4′-dimethoxytrityl)-4-N-benzoyl-3′-O-[(R,S)-N,N- diisopropylamino-(n-octyl)-phosphonyl]-2′-deoxycytidine was synthesised according to standard procedures, dissolved in dry acetonitrile (0.1 M solution) and centrifuged prior to use. Controlled pore glass supports loaded with 5′-DMT-2′-deoxyribo- nucleosides (500 Å, 100 µmol/g load) were obtained from Biosolve. The syntheses of both modified and unmodified sequences were performed on the 10 µmol scale, under standard conditions, with final deprotection of the 5′-terminal dimethoxytrityl group (trityl-off mode). The modified oligonucleotide was obtained as a mixture of two diastereoisomers due to the chirality of the n-octyl-phosphonate modification. After synthesis completion, cleavage from the resin and deblocking of the nucleobase protecting groups were achieved by treatment with a solution of saturated ammonia in MeOH (modified oligonucleotide) or with a solution of 25% ammonia in water (complementary strand) for 16 h at 50°C.
The complementary strand was purified on a Q-Sepharose anion exchange column (Pharmacia) mounted on a FPLC (Pharmacia) using a flow rate of 4 ml/min, an appropriate gradient of eluent B in eluent A and UV detection at 280 nm. Eluent A, 0.01 M NaOH; eluent B, 1.2 M NaCl in 0.01 M NaOH.
The modified oligonucleotide was first purified as described for the complementary strand, with the following adjustments: eluent A, 8:2 50 mM NH4HCO3:CH3CN; eluent B, 8:2 1 M NaCl in 50 mM NH4HCO3:CH3CN. After purification, the oligonucleotides were desalted on Sephadex G-25 columns (Pharmacia Fine Chemicals) using 0.15 M NH4HCO3 as the eluent. Separation of the two diastereomeric oligonucleotides was achieved by reverse phase HPLC on a LiChrosphere RP18e column (10 × 250 mm, 5 µm particle size; Merck) attached to a BioCAD Vision workstation (PerSeptive Biosystems). An appropriate gradient of CH3CN in 50 mM triethylammonium acetate was used as the eluent, with a flow rate of 5 ml/min and detection at 254 nm. The stereochemical configuration of the n-octyl phosphonate was S for the first eluted diastereoisomer, R for the second (see NMR assignment). After separation, both diastereoisomers were desalted on Sephadex G-25 columns as previously described.
Finally, the oligonucleotides were transformed to the corresponding Na+ form by elution on a Dowex 50Wx4 column (Na+ form) and lyophilized.
The purity of the oligonucleotides was confirmed by anion exchange chromatography and HPLC. The masses of the oligonucleotides were determined by MALDI-TOF: first eluted diastereoisomer 5′-d(CCACC*GGAAC)-3′, expected 3078.221, found 3079.1; second eluted diastereoisomer 5′-d(CCACC*GGAAC)-3′, expected 3078.221, found 3075.7; 5′-d(GTTCCGGTGG)-3′, expected 3075.034, found 3075.2.
Sample preparation
The duplex of the two stereochemically pure samples for NMR experiments were prepared by combining equimolar amounts of the alkyl-modified oligonucleotide and its complementary strand in phosphate buffer. The stoichiometry for 1:1 duplex formation was determined by performing a titration of one strand with the other. Duplex formation was monitored by UV spectroscopy. Samples were suspended in 500 µl of either D2O or 9:1 H2O/D2O. The resulting buffer solution (10 mM phosphate, 100 mM NaCl, pH 7.0) was ∼2 mM in duplex.
NMR spectroscopy
For all samples (R, S and the unmodified control duplex), all the experiments were carried out using the same experimental conditions. NMR spectra were acquired in a Bruker AMX spectrometer operating at 600 MHz and processed with the UXNMR software. DQF-COSY, hetero-COSY (1H-31P), TOCSY and NOESY experiments were recorded in D2O. The NOESY (24) spectra were acquired with mixing times of 100 and 250 ms and the TOCSY (25) spectra were recorded with the standard MLEV-17 spin-lock sequence and a 80 ms mixing time. The NOESY spectra in H2O were acquired with a 200 ms mixing time. In the experiments in D2O presaturation was used to suppress the residual H2O signal. A jump-and-return pulse sequence (26) was employed to observe the rapidly exchanging protons in 1D H2O experiments. In 2D experiments in H2O water suppression was achieved by including a WATERGATE (27) module in the pulse sequence prior to acquisition. The experiments in D2O were carried out at 25°C, below the melting temperature of the duplexes. Spectra in H2O were recorded at 5°C, to reduce the exchange with water.
The spectral analysis program XEASY (28) was used for semi-automatic assignment of the NOESY cross-peaks. Quan- titative evaluation of the NOESY cross-peak intensities was carried out automatically with the integration routines included in the package. In the case of overlapping or very weak peaks, the region of integration was selected manually. When the corresponding peaks on both sides of the diagonal were equally reliable, the average intensity was considered.
Experimental constraints
Quantitative distance constraints were obtained from NOESY experiments using complete relaxation matrix analysis with the program MARDIGRAS (29,30). A global isotropic correlation time was considered in the MARDIGRAS calculation, except for methyl groups, where a three-state jump model was used. To avoid any possible bias from the initial conditions and to estimate the error bounds in the interprotonic distances properly, different MARDIGRAS calculations were carried out with different initial models, mixing times and correlation times. Standard A- and B-form duplexes were used as initial models and three correlation times (1.0, 2.0 and 4.0 ns) were employed. Experimental intensities were recorded at two different mixing times (100 and 250 ms) for non-exchangeable protons and at a single mixing time (200 ms) for labile protons. Final distances with their errors were determined by averaging the upper and lower bounds in all individual runs. These constraints were incorporated into the AMBER potential energy by defining a flat-well potential term between the average upper and lower limits. A lower limit of 1.8 Å was set
Nucleic Acids Research, 2001, Vol. 29, No. 14 2975
for those distances where no quantitative analysis could be carried out, such as very weak intensities or cross-peaks involving labile protons. A loose distance limit of 5 Å was set for these constraints.
Sums of J-coupling constants involving H1′, H2′ and H2″ protons were estimated from DQF-COSY cross-peaks. The population of major S conformers were calculated from the sums of J-coupling constants, Σ1′ and Σ2″, according to the expressions:
fs = (31.5 – Σ2″)/10.9 and
fs = (Σ1′ – 9.8)/5.9
where fs is the population of S-type puckering (31). In those deoxyriboses where the population of the S-type conformer is >50%, torsion angle constraints were included. Since only the sums of coupling constants were estimated, loose values were set for the dihedral angle of the deoxyriboses (δ angle between 110 and 170°, ν1 between 5 and 65° and ν2 between –65 and –50°).
In addition to these experimentally derived constraints, Watson–Crick hydrogen bond restraints were used. Target values for distances and angles related to hydrogen bonds were set as described for crystallographic data (16). No backbone angle constraints were employed. Distance constraints with their corresponding error bounds were incorporated into the AMBER potential energy by defining a flat-well potential term.
Structure determination
Starting models of px DNA were built in the A and B canonical structures (32) using the INSIGHT graphics program (33). Structures were minimized in the gas phase for 5000 cycles, introducing hydrated sodium ions as counterions. Five A- and five B-form starting models for MD simulations were gener- ated by assignation of random velocities (T 300 K) to the A- and B-optimized structures. The 10 starting models were then subjected to 5 ps of MD simulation (T 300 K) for equilibration. The temperature was then gradually raised from 300 to 1000 K over 10 ps and the trajectory was followed for a further 20 ps at 1000 K. These heated structures were then cooled to 300 K over 20 ps and further equilibrated at 300 K for 20 ps. Finally, the 10 structures obtained at the end of the simulated annealing process were optimized for 8000 cycles. All these simulations were carried out using SHAKE (34) to constrain chemical bonds, using a 1 fs time step for integration. NMR constraints were used in all the simulations, as explained above.
The 10 structures (for the unmodified DNA and each diastereoisomer) obtained after the annealing process were further refined using state of the art (NMR-restrained) MD simulations considering explicit water and ions and introducing long- range electrostatic effects using the PME method (35). This lengthy procedure allows us to relax a few of the local distortions in the structure of the DNA which appear in structures derived from the simulated annealing process and that seem to be related to the neglect of solvation effects. Thus, the 10 structures obtained after simulated annealing in the gas phase were introduced in a box containing ∼2500 TIP3P (36) water molecules and sodium counterions to maintain electro- neutrality. The systems were then minimized using our standard eight-step equilibration protocol (37,38), which includes minimization and up to 160 ps of restrained MD
simulation. The equilibrated structures were then subjected to 0.5 ns of MD simulations at constant pressure (1 atm) and temperature (300 K). Periodic boundary conditions and the PME technique have been used to account for long-range effects. SHAKE (34) was used, which allowed us to use a 2 fs time step for integration. The simulations reported here represent a total of almost 20 ns of restrained PME-MD simulation in solution. This is, to our knowledge, the largest set of NMR-restrained MD simulations published. The use of very extensive MD simulations allowed us to obtain a vast sampling of the configurational space of the duplexes consistent with experimental restraints.
The AMBER-95 force field (39) was used to describe the DNA, while the TIP3P model was used to describe water molecules. The PAPQMD (40) strategy was used to para- metize the P-C torsion using HF/6-31G(d) profiles as the refer- ence. The RESP strategy (41) was used to obtain electrostatic charges for the modified oligonucleotide using HF/6-31G(d) molecular electrostatic potentials.
Analysis of structures and trajectories
Structural analyses of trajectories in solution were performed using the analysis modules in AMBER as well as in-house programs. The average structures were determined from the last 250 ps of every trajectory in solution (37,38,42). Due to the excellent convergence between the different trajectories (see below), global average structures were determined using the last 250 ps of the 10 trajectories for each diastereoisomer and the unmodified duplex. Solvation analysis was carried out using the joint ensemble of trajectories for each of the two diastereoisomers as noted elsewhere (37,38,42). Reactivity analysis was performed using the molecular interaction potential (MIP) (37,38,42), using global averaged structures. Helical analysis of MD averaged structures was carried out with the programs Curves (43) and MOLMOL (44). Reliability factors (R factors) (45) were calculated with the program CORMA (29). Solvation and MIP maps were determined using the procedure noted elsewhere (37,38,42).
RESULTS AND DISCUSSION
The numbering scheme of the duplex used in the following discussion is:
1 2 3 4 5 6 7 8 9 10 5′-d( C C A C CpxG G A A C)-3′ 3′-d( G G T G G C C T T G)-5′
20 19 18 17 16 15 14 13 12 11
where px indicates the alkyl-modified phosphonate moiety. Molecules containing each of the two possible diastereoisomers (S or R) are designated S duplex and R duplex.
Assignment
Non-exchangeable protons. Non-exchangeable protons were assigned using established techniques for right-handed, double-stranded nucleic acids using DQF-COSY, TOCSY and 2D NOESY spectra (46,47). Spin systems in the sugar moiety of the DNA were identified in the DQF-COSY and TOCSY spectra. The different intensities of the cross-peaks of H1′ with H2′ and H2″ in the NOESY spectra at short mixing times was used to stereospecifically assign H2′ and H2″. The spin systems
where connected with their own base and their 5′-neigbor in the NOESY spectra. These assignment pathways could be followed in the base–H1′ (Fig. 1) and in the base–H2′/H2″ region, for both strands. The chemical shifts of the protons in both diastereoisomers (S and R) and in the control duplex are given in Supplementary Material.
Alkyl group. Proton signals of the alkyl group in the modified strand could be assigned by combining heteronuclear and homonuclear COSY experiments. The nearest protons to the phosphonate were identified by their strong cross-peaks with the phosphorous atom in the 31P-1H hetero-COSY experiment and by the NOEs with several DNA protons (Fig. 2). The rest of the alkyl chain could be identified in the homonuclear correlation experiments, but only the protons close to the phosphorous atom could be unambiguously assigned.
Identification of the chirality. Cross-peaks between protons of the alkyl group and the DNA were identified in the NOESY spectra. The pattern of these cross-peaks is different in each sample. In one of them, strong NOEs were observed between the first two protons of the alkyl group and H8 of G6 (cross- peaks a and b in the top left panel of Fig. 2). These cross-peaks are not observed in the other diastereoisomer (see bottom right panel). Additional alkyl–DNA contacts are observed with the H3′ protons of C5 and G6 or with the H5′/5″ of G6 (cross- peaks c and f in the top right panel of Fig. 2). All possible distances between alkyl and DNA protons were checked in B-like models of the two diastereoisomers. Independently of the conformation of the alkyl chain, short distances between the first protons and H8 of G6 were found only in the S diastereoisomer. In this case, the distance to H3′ of G6 is also short, whereas the distance to H5′ of G6 is only short in the R diastereoisomer. These short distances are indicated in the molecular
models shown in the Figure 2 and correspond with the distinct cross-peak patterns observed in the NOESY fragments displayed in this figure. Therefore, the chirality of both samples could be identified unambiguously.
Exchangeable protons. Exchangeable protons were assigned from the NOESY spectra recorded in H2O (see Fig. S3 in Supplementary Material). Imino H3 protons in thymines were identified by their strong inter-strand cross-peaks with H2 of the base paired adenines. Additional cross-peaks with these H3 protons were used to identify the amino protons of adenines. Labile protons in GC base pairs were assigned by following their connection to H5 of cytosines (H5C→HN4C→H1G). NOE connectivities between imino protons of AT and adjacent GC base pairs permitted the sequential assignment of thymine H3 protons and adenine amino protons. All the labile protons were assigned, except the amino resonances of guanines. The cross-peak patterns observed for the exchangeable protons indicate that all bases form Watson–Crick pairs throughout the duplex.
Melting
The melting behavior of the two diastereoisomers is very similar. 1D spectra of the non-exchangeable protons at different temperatures are shown in Supplementary Material. No changes in the spectra were observed in either duplex below 45°C, while significant signal broadening was observed between 50 and 60°C. Melting temperatures were estimated by following the changes in thymine methyl resonances (data not shown). Under the experimental conditions used in the NMR experiments the two diastereoisomers melt at temperatures of ∼56–58°C, while the control duplex melts at ∼62°C.
Distance constraints
Quantitative distance constraints were obtained from NOESY experiments using a complete relaxation matrix analysis with the program MARDIGRAS. The total numbers of experimental distance constraints were 271 and 280 for the S and R duplexes, respectively. Intra-residual constraints involving protons of the same deoxyribose were not included, with the exception of H1′-H4′ and H2″-H4′, since they are not struc- turally relevant. Considering these constraints, the average number per base pair is almost 30, including both exchangeable and non-exchangeable protons. Distance constraint distributions for both duplexes are shown in Figure 3 and Table S5.
J-coupling analysis
J-coupling constants for deoxyriboses were estimated from DQF-COSY experiments. The sums of J-couplings for H1′, H2′ and H2″ are shown in Supplementary Material. In most cases the J-couplings are consistent with sugar puckers predominantly in the S domain. The population of the major S conformer, estimated from the sum of J-coupling constants, Σ1′ and Σ2″ (31), is >75%, except in the terminal nucleotides. The coupling constants of deoxyriboses 5 and 16 in the S duplex could not be estimated. In the cases of C5, no cross- peaks were observed in the DQF-COSY spectrum, probably due the wide line width of the H1′ signal. In the case of G16, the H2′ and H2″ resonances are degenerate.
Figure 1. H1′–aromatic region of the NOESY spectra of d(CCACCpxG- GAAC)·(GTTCCGGTGG) in D2O (100 mM NaCl, T 25°C, pH 7). Spectra of both diastereoisomers are superimposed. The R duplex spectrum is indicated by solid contours and the S duplex by dashed ones. Assignment pathways are shown for the R duplex. Intra-residual H1′–H6/H8 are labeled with the residue number (in italic for the S duplex).
Figure 2. Fragments of NOESY spectra in D2O. (Left) H2′/2″–aromatic region; (right) H2′/2″–H3′ and H2/2″–H4′/5′ region. The spectra of the S and R duplexes are shown in the upper and lower panels, respectively. Stereoviews of the C5–px–G6 regions of both duplexes are also shown. Some short inter-protonic distances, distinctive for each diastereoisomer, are indicated.
Differences between diastereosiomers
Except in the region close to the modified phosphate, both diastereoisomers present very similar chemical shifts to those of the unmodified duplexes. The chemical shifts of the exchangeable protons are very similar for the unmodified, R- and S-substituted DNAs, indicating that the alkyl group does not substantially distort the base pairs. However, significant differences are observed in the chemical shifts of non- exchangeable protons located in the neighborhood of the alkyl group. As can be seen in Figure 4, the main differences are observed in nucleotides 4, 5 and 6 and, for the S duplex, 16 and 17. Interestingly, in the case of the S diastereoisomer, the effect of the modification is observed not only in the modified strand, but also in the complementary one. The chemical shift profiles shown in Figure 4 suggest that the structure of the S duplex is more distorted, with respect to the unmodified DNA, than the R diastereoisomer.
Structure calculation
Distance and torsion angle constraints were used to calculate the structure of both modified duplexes (and the control
duplex) by restrained molecular dynamics. Two different calculations were carried out with the AMBER package (48). In the first refinement, 10 structures for each diastereoisomer (and unmodifed duplex) were calculated starting from the standard A and B models (32), with different initial velocities. These calculations were carried out in vacuo, following the high temperature annealing protocol described in Materials and Methods. Five different sets of initial velocities for each of the starting structures were used. The average value of the mutual RMS deviations between the 10 final structures was 1.6 Å, which indicates that reasonable convergence had been reached after the annealing procedure. As shown in Table 1, final distance restraint violations and ENOE terms were small, showing the ability of structures to fulfill experimental constrains. Also, the NMR R factors confirm the ability of the structures to fit experimental data (see Table 1).
Although most of the nucleic acid structures solved by NMR have been calculated using restrained molecular dynamics in vacuo, it has recently been shown that inclusion of explicit solvent terms and a better evaluation of the electrostatic term improves the quality of the final NMR-derived structures (49). We refined every final structure from in vacuo calculations by means of 0.5 ns NMR-restrained molecular dynamics trajectories, including explicit solvent terms, as described in Materials and Methods. The final MD averaged structures fit the experimental data very well, exhibiting similar figures of merit to the structures refined in vacuo (see Table 1). The structures obtained in solution are, however, better defined, with lower RMS deviation values (∼1.1 Å RMSD between the different MD averaged structures of a given diastereoisomer; see Table 2). Furthermore, local, artifactual distortions (mostly related to narrowing of the minor groove in regions distant from the lesion site) observed in the phosphate backbone of the structures obtained in vacuo are eliminated when the solvent is included in the calculations (a detailed comparison of solvated and in vacuo simulations will be provided elsewhere).
The RMSD values between the trajectories (in water) and their corresponding MD averaged conformations are 0.6–0.8 Å, indicating good stability. The RMSD between the different trajectories in water and the final R and S structures (obtained by averaging all the trajectories for each diastereoisomer) are ∼0.9–1.2 Å, demonstrating excellent convergence of all the trajectories for each diastereoisomer stability (values of ∼0.5 Å were observed for the unmodified duplex). This convergence is confirmed by the small values of the RMSD between the MD averaged structures of the different trajectories (see Tables 1 and 2). It is clear that the specific protocol used here is sufficiently effective to overcome potential barriers and any trace of the starting structure has been removed, leading to well- defined averaged structures. These structures are stable and agree very well with all the experimental restraints derived from the NMR spectra (see Table 1).
The 10 resulting structures of the final refinement in water are displayed in Figure 5 for both diastereoisomers. As can be seen in Figure 5 (and also in Table 2), the two duplexes are well defined. In general, the S duplex shows smaller ENOE and slightly smaller fluctuations with respect to the global averaged structure, which might suggest that, due to the network of distance restraints, a more clearly defined structure was obtained for this diastereoisomer. As usual in NMR-derived structures, the bases are better defined than the sugar–phosphate
Figure 3. Distribution of distance constraints for the two diastereoisomers. Bold numerals indicate the sequence order. Plain numerals indicate the number of intra-residue, sequential and inter-strand distance constraints.
backbone. Both modified and unmodified strands showed similar RMSD, indicating that the presence of the alkyl group does not introduce a dramatic change in flexibility in the modified strand.
The structure of the control duplex was also calculated following the same protocols as for the modified duplexes. Details on these calculations will be given in a forthcoming publication, where the structure of the control duplex will be fully described. In this paper we will focus only on those structural features relevant for comparison with the modified duplexes.
Description of the structures
The solution structures of the R and S duplexes belong to the B-form DNA family. The RMSD values with respect to a canonical B-type duplex are 2.2–3 Å, whereas the structure is far from the canonical A-type duplex (RMSD ≈ 4.2 Å). The RMSD of the average structure of the control duplex is 1.5–1.6 Å, which indicates that the alkyl group induces only moderate distortions in the global structure of the duplex. As expected, all the bases are in the anti orientation about the glycosidic bond and the sugars are on average ∼80% in the south (225 > P > 135) and ∼19% in the east (134 > P > 45) conformation. West puckerings (315 > P > 225) are found ∼0.9% of the time and north puckerings (45 > P > 315) are detected in <0.1% of the structures sampled during the restrained MD simulations in water. Not surprisingly, the largest deviations from south puckering are found at the transition site (C5–G6 and C15–G16). Thus, for the R diastereoisomer the deoxyribose of C15 remains in the south conformation only 45% of the time, while for the S diastereoisomer the deoxyriboses of C5 and G16 are in the south conformation only ∼30 and 59% of the time.
The helical parameters shown in Figure 6 are those expected for a B-type duplex. The only noticeable distortions are located in the base pair step of the modification. In both cases, a low rise value and high propeller twist is observed, indicating that
Table 1. Summary of NMR structure calculations
aStandard deviations in global averages (averages of the 10 MD averaged structures for each diastereoisomer). All RMSD and violation data are in Å, NOE energies are in kcal/mol.
S duplex (in vacuo)
R duplex (in vacuo)
RMSD
all heavy atomsa 1.6 ± 0.2 1.7 ± 0.4 1.1 ± 0.3 1.2 ± 0.3
backbone 1.5 ± 0.3 1.5 ± 0.4 0.9 ± 0.2 0.9 ± 0.3
all bases 1.0 ± 0.2 1.0 ± 0.3 0.7 ± 0.2 0.7 ± 0.2
Sum of violations (average)
15.5 23.6 15.7 23.7
Sum of violations (range)
Maximum violation (average)
R factor (×100) 8.6 9.5 8.4 9.3
Average NOE energy 70.1 125.0 70.4 122.6
Range of NOE energies
67.0–74 119.1–129.0 67.2–74.1 116.8–126.8
Figure 4. Chemical shift differences between non-exchangeable protons of the R and S diastereoisomers and the control duplex (plus, H6/H8; asterisk, H5/H2/Met; square, H1′; triangle, H2′; cross, H2″; circle, H3′; inverted triangle, H4′).
some slight displacement to the A-form could be induced by the presence of the alkyl phosphonate. The relatively low values of helical twist and the slight change in sugar pucker in this region may support this hypothesis. A similar effect has been observed in a DNA duplex with a phosphoramidate moiety (50), where the transition to the A-form is more dramatic.
The global RMS deviation between the two diastereoisomers is well above the RMSD of each ensemble. For instance, the average RMSD between the final S isomer structure and the 10 R trajectories is ∼1.5 Å and the RMSD between the final R isomer structure and the 10 S trajectories is ∼1.7 Å. These
values are ∼0.4–0.7 Å larger than those found when the different trajectories of one diastereoisomer are compared with the final structure of the corresponding diastereoisomer and ∼0.8–1 Å larger than those obtained when the trajectories are compared with their own averaged structure. These findings clearly indicate that the structural differences between both diastereoisomers are small, but not negligible. Analysis of local geometrical parameters shows that the important changes are located in the modified phosphate and the surrounding area. As expected from chemical shift data (Fig. 4), larger differences in phase and glycosidic angles (Fig. 7) between the two diastereoisomers are found only for residues 5, 6, 15 and 16. Backbone torsion angles also show different values, but these torsion angles are not so well defined in the final ensem- bles of each diastereoisomer and the differences cannot be considered meaningful.
The presence of a neutral alkyl phosphonate unit in the middle of the helix induces moderate bending in the DNA, as is clearly shown in Figure 8. The DNA bends towards the major groove, whose width (defined as the P–P distance) is reduced by ∼2 (R) and 3 (S) Å from canonical B-DNA structures, leading to the generation of a global bending in the helix, as shown in a shortening of 2.6 (R) and 3.6% (S). This bending is almost negligible in the control duplex (see Fig. 8), where the shortening is only 0.9%. The capability of NMR methods to detect bending in short DNA fragments has been discussed for a long time (51,52). The experimental information observed by classical NMR methods (NOEs and J-couplings) is mainly local and, consequently, the local geometry is better defined than the global shape of the molecule. Even high resolution DNA structures, obtained with a large number of very accurate experimental constraints, may not be sufficiently defined to determine global parameters, such as bending, with precision. A recent approach, based on the measurement of residual dipolar couplings in partially oriented nucleic acids,
Figure 5. Superposition of 10 final MD averaged structures for the R and S duplexes.
Table 2. RMSDs (in Å) between the 10 final structures and the two initial A and B canonical models
Upper right is the R duplex, lower left is the S duplex.
1 2 3 4 5 6 7 8 9 10 A B
1 2.18 1.63 1.67 2.03 1.77 1.43 1.64 1.37 1.60 3.98 3.54
2 1.03 1.47 2.32 1.84 2.17 2.09 2.21 2.21 2.01 4.72 2.95
3 2.02 2.27 1.84 1.64 1.66 1.65 1.65 1.43 1.66 4.49 2.92
4 1.83 2.13 1.04 1.61 1.60 1.47 2.04 1.98 1.64 4.27 2.86
5 1.18 1.29 1.75 1.59 1.69 1.67 1.92 1.94 1.47 4.79 2.50
6 2.39 2.59 2.16 2.19 2.16 1.09 1.79 1.63 1.16 4.72 2.64
7 1.78 1.71 1.66 1.67 1.68 2.16 1.46 1.33 1.02 4.24 2.85
8 1.90 2.16 2.24 2.28 1.97 1.26 1.97 0.92 1.56 4.63 3.15
9 1.41 1.48 1.92 1.77 1.39 1.76 1.30 1.55 1.40 4.43 3.20
10 1.74 2.04 1.68 1.66 1.62 1.61 1.66 1.61 1.28 4.54 2.50
A 4.67 4.89 4.30 4.26 4.49 3.68 4.46 3.90 4.18 4.25 5.81
B 2.21 2.20 3.17 3.11 2.53 3.29 2.85 2.86 2.49 2.58 5.81
has proved useful in determining global geometric parameters (53). Unfortunately, such an approach cannot be used in this case, since it requires samples isotopically labelled with 13C and 15N. On the other hand, state of the art molecular dynamics methods can be of great help in improving the quality of NMR structures (49). In fact, unrestrained molecular dynamics simulations have been able to reproduce both global and local properties of DNA with enough fidelity (54).
To determine the significance of the average bending observed in the S and R duplexes, we have carried out an error analysis with the ensemble of averaged structures obtained for
the 10 different trajectories in water. As expected, the standard deviations for shortening are rather large for both stereoisomers, ∼0.8 and 1.2% for the R and S duplexes, respectively, which suggests that global bending movements should be one of the principal components of the essential dynamics of the modified duplexes (as a comparison, the standard error for shortening of the unmodified duplex is only 0.2%). These deviations represent ∼23 and 33% of the total shortening found in the modified duplexes. Accordingly, despite the large error bars, the observed bending must be considered meaningful.
Interestingly, the S duplex, where the alkyl group is pointing into the helix, leads to the largest bending of the helix. This indicates that although an electrostatic effect is probably the main driving force for DNA bending, steric effects are also important. This result agrees with gel migration experiments carried out with methylphosphonates (12,13). In such experiments the bending observed in a racemic sample was larger than in samples with the pure (Rp) stereoisomers. The structures reported in this paper are direct evidence that S stereoisomers produce a larger bending than R stereoisomers. In our case, this difference was probably enhanced because of the larger size of the alkyl chain in the modified phosphate.
Flexibility
In general both stereoisomers are equally well defined. However, our results indicate that the S duplex is slightly more flexible than the R duplex in the neighborhood of the modified phosphate. Furthermore, analysis of RMSD fluctuations between the trajectories and the global MD averaged structures (see above) suggest that both modified duplexes are more flexible than the control DNA. The lack of H1′-H2′ and H1′-H2″ COSY cross-peaks for C5 and the chemical shift degeneration between H2′ and H2″ of G16 indicate that these two sugars are involved in a dynamic process. Although the J-coupling constants for the protons of these sugars could not be estimated, it is reasonable to assume that this effect is caused by a large population of no-S puckerings for these two deoxyriboses. The 10 refined conformers of the S duplex present sugar puckers of C5 and G16 in the S region, although large standard deviations for pseudorotation phase angles are observed for these residues (Fig. 7), confirming the larger flexibility of sugars in this part of the helix for the S diastereoisomer. Analysis of the sugar puckering along the trajectory (see above) shows that the deoxyriboses of these residues are especially flexible for the S diastereoisomer. Thus, the deoxyribose of G16 of the S diastereoisomer is 59% of the time in the south region, 25% in the east, 15% in the west and 1% in the north region. Similar flexibility is shown by the deoxyribose of C5 of the S diastereoisomer, which is 68% of the time in the east, 31% in the south and 1% in the west regions. The sugars of the R diastereoisomer are more rigid in this region (80–90% of the population in the south area), except for the deoxyribose of C15, which is 54% of the time in the east region and ∼44% in the south region. In the control duplex, the cross-peak pattern in the DQF-COSY experiment indicates that all the non-terminal deoxyriboses are predominantly in the south conformation. Therefore, the enhanced flexibility observed in the modified duplexes must be caused by the presence of the alkyl group.
Figure 6. Helical parameters for the averaged structures of the R (solid squares) and S duplexes (open circles and dashed lines). Error bars are the RMSD of the helical parameters between the 10 refined structures of each diastereoisomer.
Molecular recognition characteristics
The analysis of the 2 (isomers) × 10 (trajectories) × 0.5 ns trajectories of the hydrated DNA systems allowed us to obtain a representation of the recognition characteristics of the R and S duplexes, which can be compared to those of the unmodified duplex. The modified DNA is well solvated and shows characteristics of hydration not too different from those of an unmodified DNA. Solvation contours obtained by integrating the water population in the different trajectories (see Fig. 9) show the existence of regions of dense hydration in the minor groove, as well as smaller regions in the major groove in d(G·C) steps, a situation typical of B-type duplexes (55). These regions are reduced surrounding the phosphonate unit. No extreme differences are evident between the two diastereoisomers, although solvation in the minor groove of the R diastereoisomer is slightly better than that of the S diastereoisomer.
MIPs provide a general view of the ability of a DNA to interact with a small cationic molecule (37,38,42). The minor groove is the most reactive region of both the unmodified and the modified DNAs (see Fig. 10), but negative regions are located around the central d(G·C) steps. The presence of the alkyl phosphonate unit produces a reduction in the reactivity of
the DNA with cations, especially in the minor groove, in good agreement with changes in the hydration pattern (see Fig. 9). This change in MIP can be easily understood considering the reduction in positive charge upon phosphate→alkyl phosphonate substitution. No large differences are found between the two diastereoisomers in terms of MIPs contours.
Biological implications
Our NMR-restrained simulations confirm previous theories that phosphate neutralization can induce DNA bending (13), even when the general characteristics of a B-DNA are preserved. The type of bending described here in microscopic detail might, however, be somehow different to that detected in long fragments of DNA when several phosphates are neutralized (13). Thus, the small magnitude of the bending found here might not be easy to detect in large fragments of DNA, where neutralization of several phosphate groups might be necessary. Since only one phosphate group is neutralized, bending should occur towards the more flexible major groove [which in d(G·C) sequences is very negatively charged (see Figs 9 and 10)] and not towards the backbone. At this point, it is remarkable to see (Figs 9 and 10) that the grooves, and not the vicinity of
Figure 7. Averaged sugar pseudorotation phase angle and glycosidic torsions for the R (solid squares) and S (open circles and dashed lines) duplexes.
Figure 8. Global MD averaged structures (obtained by averaging the 10 MD averaged structures) of the R and S duplexes. The helical axis is displayed to show the magnitude of bending. The MD averaged structure of the control duplex with the helical axis is displayed for comparison.
the phosphates, concentrate most of the negative charge of the DNA, which explains the finding that the neutralization- induced bending found in our structure is groove-driven and not backbone-driven. On the other hand, the fact that both the R and S stereoisomers show detectable bending indicates that steric effects can be important in determining bending, as noted by the difference in bending of the R and S stereoisomers, but it is not the driving force. Our high resolution structures support the theory that ‘in-phase’ neutralization of the major groove of DNA induced by histones of the nucleosome could be the driving force responsible for bending of DNA around nucleosomes. It is worth noting that the bending of DNA does not introduce dramatic changes in the general structure of DNA, in agreement with the known structure of DNA bound to nucleosomes. It is suggested that neutralization-induced bending is mostly a local distortion of DNA, which does not have long- range effects on the physiological activity of the DNA.
The results reported here provide clear support for the idea that simple neutralization of one phosphate can induce a non- negligible bending in the DNA structure. Accordingly, the driving force for bending found in many DNA–protein complexes (16–20) might be asymmetrical neutralization of phosphates by the cationic environment of DNA-binding proteins, rather than specific hydrogen bonding interactions in the grooves.
SUPPLEMENTARY MATERIAL
ACKNOWLEDGEMENTS
This work was supported by the EU Biotechnology Program (BIO-CT96-0509), the Spanish DGICYT (PB98-1222, PM99- 0046 and PB97-0941-C02-02), the Wellcome Trust and the
Figure 9. Preferential regions of hydration for the R, S and control duplexes. Regions in red correspond to a density of 4 g/ml.
Figure 10. MIP maps for the R, S and control duplexes. Regions in red correspond to interaction energies of –5 kcal/mol. O+ was used as the probe molecule.
Centre de Supercomputació de Catalunya (Molecular Recogni- tion Project).
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