Sequence-dependent nucleotide dynamics revealed by intercalated ring r
http://www.100md.com
《核酸研究医学期刊》
MRC Laboratory of Molecular Biology, Hills Road, Cambridge CB2 2QH, UK
* To whom correspondence should be addressed at present address: Medivir UK Ltd, 100 Fulbourn Road, Cambridge CB1 9PT, UK. Email: Jose.Gallego@medivir.com
ABSTRACT
Bisnaphthalimide intercalators are anti-tumour agents composed of two planar rings linked by a flexible diazanonylene chain. The intercalated rings of three bisnaphthalimide analogues complexed to DNA are found here to undergo 180° rotating motions that do not affect the diazanonylene linker atoms bound to the major groove. These ring rotations are detected by NMR spectroscopy in a broad range of sequence contexts and duplex lengths. A comparative analysis of the frequency and activation energies of such excited states in different complexes and conditions indicates that these motions (i) are unrelated to drug dissociation; (ii) are a consequence of concerted, sequence-dependent nucleotide movements taking place on the millisecond time scale; and (iii) may occur inside the DNA duplexes. The rotation frequencies range from 2 to 25 s–1 at 25°C, depending on DNA composition and the size of the rotating rings. The detected nucleotide dynamics are likely to play an important role in the binding kinetics of the numerous proteins and drugs that require base unstacking when interacting with DNA.
INTRODUCTION
Nucleic acid dynamics influence the binding kinetics of transcription factors (1,2), endonucleases (3,4) and drugs (5–9) that intercalate or thread through the DNA double helix, and are important in DNA processing events requiring base flipping from the double helix (10,11) and strand stretching (12). Functionally relevant nucleotide motions have also been reported in ribozymes (13) and ribosome subunits (14,15). Whereas there is essential structural insight into most of these processes, a significantly smaller quantity of data is available about the direction and extent of nucleotide motions. This is especially true for time scales that are biologically (2,11,13,14,16) and pharmacologically (5–7) relevant but currently inaccessible to molecular dynamics simulations.
We recently solved the solution structure of a complex between elinafide (LU) (Figure 1a) and d(ATGCAT)2 (d6) (17). Using NMR spectroscopy, we found that the two naphthalimide rings of the drug bis-intercalate at the TpG (CpA) steps of d6, and undergo 180° rotations without affecting the 3,7-diazanonylene linker atoms bound to the DNA major groove (17) (Figure 1b). Here I have analysed these rotating motions in complexes of LU, LU-84743 (LUNH) and LU-77655 (LUNO) (18) (Figure 1a) with DNA duplexes of varying length and sequence (Figure 1c) and under different temperature and solution conditions. A comparative analysis of these complexes indicates that the rotational motion of the naphthalimide rings is detected in all sequences, and reveals sequence-dependent DNA nucleotide movements taking place on the millisecond time scale.
Figure 1. Bisnaphthalimide intercalators and DNA–bisnaphthalimide complexes. (a) Chemical structure of bisnaphthalimides. The van der Waals diameters (d) of the rotating naphthalimide rings are specified. (b) Schematic representation of a DNA–bisnaphthalimide complex and of the observed rotation of the naphthalimide rings (grey rectangles). The activated state is shown in a darker tone, and ring rotation may take place outside or inside the intercalation site. (c) The DNA–bisnaphthalimide complexes analysed in this study. The DNA sequences are self-complementary and form antiparallel duplexes containing 6 to 42 base pairs. The bisnaphthalimide molecules are shown schematically.
MATERIALS AND METHODS
Sample preparation
The DNA–bisnaphthalimide complexes were prepared as described (17) after purification of the DNA oligomers by reverse-phase high-performance liquid chromatography (HPLC) or gel electrophoresis. All samples contained 1 mM complex in 20 mM sodium phosphate buffer (pH 7), 0.2 mM EDTA, and 0 or 150 mM NaCl.
NMR spectroscopy
All NMR spectra were acquired on temperature-calibrated Bruker DRX-500 and DMX-600 spectrometers and processed with Felix 97.0 software. Each complex was analysed at several temperatures using 1H2O NOESY and a series of 2H2O NOESYs uninterruptedly collected at several mixing times (typically 0, 5, 10, 15, 20, 40, 80 and 200 ms), together with 2H2O pure exchange (19), dqf-COSY, TOCSY and ROESY experiments, all with recycle delays of 2 s. All LU, LUNH and LUNO protons and most of the aromatic, H1', H2' and H2'' DNA resonances were assigned in each of the complexes and in each of the unbound DNA duplexes studied. All sequences form the self-complementary duplexes shown in Figure 1c, although a minor hairpin form was also detected for d26.
LU ring rotation rate constants were measured at an average of nine temperatures (ranging from –6 to 56°C) in each of the eleven DNA–LU complexes and ionic strength conditions studied (Table 1; Figure 3). Depending primarily on temperature and complex size, the rates were determined from a combination of NOESY experiments, magnetization transfer data and lineshape analyses. The NOESY and magnetization transfer methods were generally applied at lower temperatures and used the H4–H9 pair of exchanging naphthalimide resonances of the LU complexes. H4 and H9 are 6.7 ? apart (Figure 1a), and the H4 resonance is shifted upfield and resolved in all complexes (Figure 2b). Magnetization transfer experiments were carried out as described before (17), at an average of two different temperatures per complex and solution condition. For NOESY measurements (on average, one per complex and solution condition), the rates were obtained using the H4 diagonal volume and the initial build-up of the H4–H9 crosspeak volume as a function of at least three mixing times, ranging from 0 to 50 ms (20). At higher temperatures, the rates were determined (at an average of six different temperatures per complex and solution condition) with the lineshape analysis program Mexico (21) using all the resolved LU naphthalimide resonances. The coupling constants of the naphthalimide system and the temperature dependence of the naphthalimide 1H chemical shifts were included in the calculations, and the reference transverse relaxation rate, T2–1, was obtained at each temperature from the linewidth of resolved aromatic DNA peaks not affected by exchange. Assuming that they are independent of temperature (Figure 3), H* and S* for LU ring rotation were obtained by fitting the rate constant (k) and temperature (T) data to the Eyring expression, ln(k/T) = ln(kb/h) + S*/R – H*/RT, where h, kb and R are Planck's constant, Boltzmann's constant and the universal gas constant, respectively. DNA–bisnaphthalimide dissociation rates were determined from NOESY experiments using the TGCA H2 resonance, shifted upfield and resolved (Figure 2b, inset) in all complexes except d22–LU, and its crosspeak to TGCA H2 in residual free DNA (Figure 2a, asterisk). All linear and non-linear (Levenberg–Marquardt) regression analyses were performed with the program Pro Fit 5.5.3. LUNH and LUNO ring rotation rates and activation parameters were determined from NOESY experiments as described for LU.
Table 1. Ring rotation rates and activation energies, dissociation rates, and thermal melting temperatures of DNA–bisnaphthalimide complexes
Figure 3. Naphthalimide ring rotation rate constants as a function of temperature and: (a) duplex length, (b) ionic strength, (c) sequence, (d) ring location and (e) ring rotating diameter. (a) d6–LU, d8–LU, d10–LU, d20–LU and d42–LU, 0 mM NaCl; (b) d10–LU and d20–LU, 0 and 150 mM NaCl; (c) d22–LU and d22a–LU, 150 mM NaCl; (d) d26–2LU, internal and external probe location, 150 mM NaCl; (e) d22a–LU, d22a–LUNH and d22a–LUNO, 150 mM NaCl. The rate constants (k) are in s–1, and the temperatures (T) in K. In (d), internal and external refer to the position of the LU rings in the d26–2LU complex (Figure 1c).
Figure 2. d20–LU 1H NMR spectra. (a) Aromatic region of the pure exchange spectrum (500 MHz, mixing time 60 ms) of d20–LU at 28°C, showing the crosspeaks between symmetrical and exchanging naphthalimide protons (Figure 1a). The exchange crosspeak identified with an asterisk connects the bound and unbound resonances of H2 of the adenine base adjacent to the naphthalimide rings in all complexes (Figure 1c). The intensity of this crosspeak is much smaller than the intensity of the crosspeaks connecting the exchanging naphthalimide ring protons, indicating that ring dissociation is slower than ring rotation. (b) 1H2O and 2H2O (inset) 1H NMR spectra (500 MHz) of d20–LU at 20 and 24°C, respectively. The LU H4 and TaHN3, TbHN3, AcH2 and GHN1 assignments of the d(A8TaGCAcTbT7)2 d20 sequence are labelled over the corresponding resonances. In (a) and (b), the axes represent 1H frequency in parts per million.
Error analysis
For NOESY and magnetization transfer experiments, the rate errors were estimated from the standard deviation of the parameters fitted by exponential or linear regression, and were within 2–6% and 5–15%, respectively. NOESY determinations included the standard deviation of the peak volumes in the calculations. For LUNH and LUNO, the average error is slightly larger (15% on average) due to smaller exchange rates. For LU lineshape analyses, the rate errors were estimated to be 10%, based on visual inspection and variation of the reference T2–1 values. The H*and S* errors (Table 1) were then obtained by propagation analysis from the standard deviations of the Eyring fits, which included the rate errors and a systematic 0.1°C temperature error in the calculations.
UV melting experiments
The thermal denaturation of the DNA–LU complexes was monitored by measuring the UV absorbance at 260 nm as a function of temperature in Beckman DU 650 or Varian Cary 500 Scan spectrophotometers. The temperature was raised to 90°C at a gradient of 0.5°C min–1 and subsequently decreased at the same rate. The concentration of DNA–LU was 0.5–1 ODU ml–1 (1–8 μM) in solutions identical to those used for NMR experiments.
RESULTS
LU ring rotation
Analysis of the NMR data confirmed that, as in d6–LU (17), the two rings of LU, LUNH and LUNO bis-intercalate into the TGCA sequence from the major groove in all complexes studied (Figure 1b). There are no indications of interaction with any other region of the oligomers. LU, LUNH and LUNO ring rotation is also observed in all complexes, as demonstrated by the presence of exchange crosspeaks between naphthalimide proton resonances (Figure 2a), detected by NOESY experiments at very short (3 ms) mixing times and confirmed by exchange experiments; the selective broadening of these naphthalimide resonances as the exchange rate increases (17); and the absence of exchange crosspeaks or broadening affecting the diazanonylene linker methylene hydrogens (17). These three observations demonstrate the occurrence of selective 180° rotations of the naphthalimide rings, which exchange between two intercalated states without affecting the linker atoms bound to the DNA major groove.
Additionally, naphthalimide ring rotation occurs at temperatures well below the melting points determined by UV and NMR thermal denaturation experiments (Table 1); the DNA base pairs are stacked and hydrogen-bonded in all complexes, as demonstrated by the observation of chemical shifts and inter-strand NOEs typical of double-helical DNA (Figure 2b); no changes in DNA or drug chemical shifts are observed at high temperatures, indicating that the complexes retain the bis-intercalated conformation near their melting points (17), and the ring rotation rates are significantly faster than the DNA–bisnaphthalimide dissociation rates (Figures 2a and 4a; Table 1).
Figure 4. (a) Comparison of d20–LU dissociation rates (blue diamonds; s–1) and d20–LU ring rotation rates (pink squares) at 150 mM NaCl and three different temperatures: 8, 16 and 24°C. Dissociation rates are less sensitive to temperature and 5- to 9-fold slower than ring rotation rates. (b) Imino proton linewidths (, Hz) as a function of temperature (T, °C) for free d10 (green lines) and d10–LU (red lines). Triangles and squares correspond to the Ta and Tb HN3 linewidths of the AAATaGCATbTT d10 sequence, respectively. The G HN1 linewidth values are connected by bare lines.
In the following sections, the frequencies and activation energies for LU, LUNH and LUNO ring rotation are analysed in different sequence, temperature and solution environments.
Duplex length
The effect of the number of flanking bases on ring rotation rates and activation energies was studied in a series of LU complexes with d6, d8, d10, d20 and d42 (Figure 1c) at 0 mM NaCl. The probe rotation rates are almost 10-fold slower in d8–LU0, with three base pairs stacked on either side of the LU rings, compared with d6–LU0, which contains only two. In contrast, the relative differences between d8–LU0, d10–LU0, d20–LU0 and d42–LU0 are smaller. In fact, the rotation rates in d42–LU0, with two full helix turns stacked on each of the LU rings, are slightly faster relative to the shorter d20–LU0 complex (Figure 3a; Table 1). The ring rotation activation enthalpies (H*) and entropies (S*) are also affected by duplex length and tend to decrease when the number of flanking base pairs increases (Table 1).
Effect of ionic strength
This was analysed by comparing d10–LU, d20–LU and d42–LU at 0 and 150 mM NaCl. Due to the screening of intermolecular electrostatic interactions, the presence of salt in the solution increases the thermal stability of the complexes, but also tends to increase the DNA–LU dissociation rates (Tm and kd, Table 1). In contrast, the rotation rates slow down significantly at 150 mM NaCl for the d10–LU system. The effect is small in the two larger complexes (Figure 3b; Table 1).
Sequence
The effect of base sequence on ring rotation was studied by analysing d22–LU and d22a–LU at 150 mM NaCl. In d22–LU the probe rings are flanked by dA10?dT10, whereas in d22a the flanking sequences are alternating, d(AT)5?d(AT)5 (Figure 1c). Homopolymeric A:T tracts adopt the relatively rigid and remarkably stable B' conformation (22), whereas alternating ones are more flexible (23) and thermodynamically less stable (24). This is reflected in the higher melting temperature of d22–LU relative to d22a–LU (Table 1). The DNA–LU dissociation rate is also slower in d22–LU than in d22a–LU (Table 1) but strikingly, the probe rotation rates are faster in the more stable d22–LU complex (Figure 3c; Table 1).
Internal versus external probe location
Since LU binds to any DNA sequence containing guanine (J. G., unpublished data), d26 was designed to also probe sequences flanked by G:C pairs (Figure 1c): two LU molecules bind to each of the TGCA sites so that, for each of them, one of the rings is in an external position, adjacent to one of the homopolymeric dA5?dT5 tracts, and the other one is in an internal position, adjacent to the alternating d(AT)6?d(AT)6 central tract. The presence of two LU molecules bound at both ends of d26 is expected to have a strong stabilizing effect, because bis-intercalating agents have been shown to clamp the nucleotides sandwiched between the drug rings and to strongly increase the lifetime of those base pairs (25). Indeed, the d26–2LU complex has the highest melting temperature (Table 1). Additionally, the different chemical environments resolved the internal and external LU H4 resonances, allowing for the comparison of the ring rotation rates at the two locations. The results are consistent with the previously observed trends: first, the internal and external rates are different, with the internally located rings rotating more slowly than the external ones (Figure 3d; Table 1); and second, the rotating motions at the internal location are favoured by a smaller H* but disfavoured by a more negative S*, whereas the opposite is true for the external rings (Table 1).
Diameter of the rotating rings
This effect was analysed by comparing d22a–LU, d22a–LUNH and d22a–LUNO at 150 mM NaCl. In contrast to LU and due to the asymmetry of their rings (Figure 1a), LUNH and LUNO give rise to two exchanging, unequally populated sets of five naphthalimide resonances (data not shown). A distinct correlation is detected between the rotation rates and the van der Waals diameter of the rotating rings (Figure 1a), with the rates slowing down markedly for LUNH and LUNO relative to the unsubstituted bisnaphthalimide (LU) (Figure 3e, Table 1).
DISCUSSION
Several conclusions can be reached from the above results. Intercalated ring rotation occurs in sequences resembling polymeric DNA and at internal locations protected from the duplex ends by base pair clamping, as can be deduced from the analysis of d6–LU, d8–LU, d10–LU, d20–LU, d42–LU and d26–2LU (Figure 3a and d; Table 1). Additionally, these rotating motions are detected at temperatures well below the UV and NMR melting points of all DNA–LU complexes (Table 1). With the possible exception of the faster rates detected in the smaller d6–LU complex, these observations rule out the possibility of ring rotation being a consequence of end effects in the DNA double helices.
A number of observations also demonstrate that intercalated ring rotation is unrelated to DNA–bisnaphthalimide dissociation. Depending on the complex and the temperature, DNA–bisnaphthalimide dissociation rates are 2- to 23-fold smaller than the ring rotation rates (Figures 2a, 4a; Table 1). Dissociation rates are also less dependent on temperature than rotation rates (Figure 4a). The d10–LU rotation rates slow down when the ionic strength is increased and the DNA duplex is stabilized (Figure 3b; Table 1, k and Tm). In contrast, the DNA–ligand dissociation rates accelerate when the ionic strength of the solution is higher (Table 1, kd), probably due to electrostatic screening of intermolecular interactions.
The observed stacked ring rotation is sequence dependent. The base sequences of d22 and d22a differ at positions that are not immediately adjacent to the naphthalimide rings, but have different conformational (22,23) and thermodynamic (24) properties. Comparison of d22–LU and d22a–LU indicates that the rotation rates reflect the sequence-dependent dynamic properties of DNA, and do not necessarily correlate with the thermal stability of the complexes or the observed DNA–ligand dissociation rates (Figure 3c; Table 1).
The rotation rate constants are affected by both the number and the sequence of flanking bases that are not adjacent to the naphthalimide rings, and depend on the internal or external location of the LU rings within the d26–2LU complex (Figure 3a, c and d; Table 1). In fact, the activation entropies (Table 1, S*) tend to become more negative as additional flanking base pairs are added or when LU ring rotation involves the internal base pairs of d26. Taken together, these observations indicate that stacked ring rotation implicates concerted base movements within the DNA–LU complexes: the probability of such concerted motions is expected to diminish with the presence of more flanking base pairs or at internal locations protected from the duplex ends by base pair clamping, hence the diminishing S* values.
Two mechanisms of intercalated ring rotation can be envisioned to explain the observations listed above. Ring rotation could take place inside the DNA duplexes, or alternatively the rings could independently slide out of the helix, flip in solution and intercalate back into DNA. The NMR experiments only provide information on the frequency and activation energies of the excited states, but not on their nature. It is therefore difficult to completely rule out one of the two possible mechanisms based on indirect evidence.
However, a number of observations suggest that ring rotation takes place inside the DNA duplexes. The external mechanism of rotation would likely involve part of the amino-alkyl linker protons, yet these protons are unaffected by naphthalimide exchange (17). Both the rates and the activation energies of LU ring rotation are exquisitely sensitive to the sequence and length of the flanking segments and to the external or internal location of the drug rings. An approximate 10-fold reduction is observed when the van der Waals diameter of the rotating rings is increased by 1 ? (Figures 1a and 3e; Table 1), and ring rotation is completely abolished in LU diimide analogues (26). A mechanism based on independent ring dissociation and rotation in the solvent could also account for the observed LU, LUNH and LUNO rate differences. However, since the three analogues share the same diazanonylene linker, one would expect a correlation between the ring dissociation/rotation frequency and the expected d22a–LU, d22a–LUNH and d22a–LUNO dissociation constants, which can be estimated from the relative Tm values of the corresponding complexes (27). This correlation is not observed (Table 1), indicating that ring rotation is unrelated to DNA dissociation.
If ring rotation takes place inside the DNA duplexes, the activated state of the rotating probes (with 9.2–10.8 ? van der Waals diameters) must require a transient interbase distance strikingly longer that the typical 3.4 ? stacking distance between bases. Stacked ring rotation may be facilitated by the spontaneous flipping (opening) of adjacent DNA base pairs in the complex. Although at significantly faster rates than the ring rotation rates reported here, base pair opening events also occur on the millisecond time scale (11,25). Ring rotation may also be assisted by spontaneous stretching and buckling nucleotide motions within the DNA duplex. Whereas the DNA base pairs are stacked and hydrogen-bonded in all complexes (Figure 2b), significant broadening of the imino proton resonances of the thymine DNA bases is observed in all sequences upon LU complex formation (Figure 4b; Supplementary Material). The observed broadening supports the occurrence of increased base motions in the bound DNA duplexes, which may facilitate in situ rotation of the intercalated rings.
The nucleotides comprising the immediate binding environment of the bisnaphthalimide ligands are the same in all complexes (TGCA), yet the ring rotation rates and activation parameters are influenced by the length, sequence and context of the adjacent nucleotide segments. Thus, irrespective of the rotation mechanism, it can be unambiguously concluded that the different naphthalimide ring rotation rates reflect the dynamic properties of each bound DNA sequence, and are facilitated by concerted nucleotide movements taking place at rates of 2–30 s–1 at 25°C. The detected dynamics are likely to play an important role in the binding kinetics of the numerous proteins and drugs that require unstacking of adjacent bases in order to intercalate or thread through the DNA double helix, and may influence DNA processing events involving separation and rejoining of complementary bases.
SUPPLEMENTARY MATERIAL
ACKNOWLEDGEMENTS
The initial stages of this work were carried out in the laboratory of Brian Reid at the Chemistry Department of the University of Washington, Seattle, USA, to whom I am grateful. I also thank Elisabeth Golden and David Loakes for help in the laboratory, Federico Gago, Miguel Bra?a and Christian Bailly for providing the bisnaphthalimide drugs, the EU Commission for a 2000–2002 Marie Curie fellowship, and the MRC for financial support.
REFERENCES
Werner,M.H., Gronenborn,A.M. and Clore,G.M. ( (1996) ) Intercalation, DNA kinking, and the control of transcription. Science, , 271, , 778–784.
Wu,J., Parkhurst,K.M., Powell,R.M., Brenowitz,M. and Parkhurst,L.J. ( (2001) ) DNA bends in TATA-binding protein–TATA complexes in solution are DNA sequence-dependent. J. Biol. Chem., , 276, , 14614–14622.
Tsutakawa,S.E., Jingami,H. and Morikawa,K. ( (1999) ) Recognition of a TG mismatch: the crystal structure of very short patch repair endonuclease in complex with a DNA duplex. Cell, , 99, , 615–623.
Horton,N.C., Dorner,L.F. and Perona,J.J. ( (2002) ) Sequence selectivity and degeneracy of a restriction endonuclease mediated by DNA intercalation. Nature Struct. Biol., , 9, , 42–47.
Fox,K.R., Brassett,C. and Waring,M.J. ( (1985) ) Kinetics of dissociation of nogalamycin from DNA: comparison with other anthracycline antibiotics. Biochim. Biophys. Acta, , 840, , 383–392.
White,R.J. and Phillips,D.R. ( (1989) ) Sequence-dependent termination of bacteriophage T7 transcription in vitro by DNA-binding drugs. Biochemistry, , 10, , 4277–4283.
Tanious,F.A., Yen,S.F. and Wilson,W.D. ( (1991) ) Kinetic and equilibrium analysis of a threading intercalation mode: DNA sequence and ion effects. Biochemistry, , 30, , 1813–1819.
Williams,H.E.L. and Searle,M.S. ( (1999) ) Structure, dynamics and hydration of the nogalamycin–d(ATGCAT)2 complex determined by NMR and molecular dynamics simulation in solution. J. Mol. Biol., , 290, , 699–716.
Onfelt,B., Lincoln,P. and Norden,B. ( (2001) ) Enantioselective DNA threading dynamics by phenazine-linked 2+ dimers. J. Am. Chem. Soc., , 123, , 3630–3637.
Roberts,R.J. and Cheng,X. ( (1998) ) Base flipping. Annu. Rev. Biochem., , 67, , 181–198.
Klimasauskas,S., Szyperski,T., Serva,S. and Wüthrich,K. ( (1998) ) Dynamic modes of the flipped-out cytosine during HhaI methyltransferase–DNA interactions in solution. EMBO J., , 17, , 317–324.
Nishinaka,T., Ito,Y., Yokoyama,S. and Shibata,T. ( (1997) ) An extended DNA structure through deoxyribose-base stacking induced by RecA protein. Proc. Natl Acad. Sci. USA, , 94, , 6623–6628.
Zhuang,X., Kim,H., Pereira,M.J., Babcock,H.P., Walter,N.G. and Chu,S. ( (2002) ) Correlating structural dynamics and function in single ribozyme molecules. Science, , 296, , 1473–1476.
Rodnina,M.V. and Wintermeyer,W. ( (2001) ) Fidelity of aminoacyl-tRNA selection on the ribosome: kinetic and structural mechanisms. Annu. Rev. Biochem., , 70, , 415–435.
Spirin,A.S. ( (2002) ) Ribosome as a molecular machine. FEBS Lett., , 514, , 2–10.
Ukiyama,E., Jancso-Radek,A., Li,B., Milos,L., Zhang,W., Phillips,N.B., Morikawa,N., King,C.Y., Chan,G., Haqq,C.M., Radek,J.T., Poulat,F., Donahoe,P.K. and Weiss,M.A. ( (2001) ) SRY and architectural gene regulation: the kinetic stability of a bent protein–DNA complex can regulate its transcriptional potency. Mol. Endocrinol., , 15, , 363–377.
Gallego,J. and Reid,B.R. ( (1999) ) Solution structure and dynamics of a complex between DNA and the antitumor bisnaphthalimide LU-79553: intercalated ring flipping on the millisecond time scale. Biochemistry, , 38, , 15104–15115.
Bailly,C., Brana,M. and Waring,M.J. ( (1996) ) Sequence-selective intercalation of antitumour bis-naphthalimides into DNA. Evidence for an approach via the major groove. Eur. J. Biochem., , 240, , 195–208.
Fejzo,J., Westler,W.M., Macura,S. and Markley,J.L. ( (1991) ) Strategies for eliminating unwanted cross-relaxation and coherence-transfer effects from two-dimensional chemical-exchange spectra. J. Magn. Reson., , 92, , 20–29.
Perrin,C.L. and Dwyer,T.J. ( (1990) ) Application of two-dimensional NMR to kinetics of chemical exchange. Chem. Rev., , 90, , 935–967.
Bain,A.D. and Duns,G.J. ( (1995) ) A new approach to the calculation of NMR lineshapes of exchanging systems. J. Magn. Reson. A, , 112, , 258–260.
Nelson,H.C.M., Finch,J.T., Luisi,B.F. and Klug,A. ( (1987) ) The structure of an oligo(dA)·oligo(dT) tract and its biological implications. Nature, , 330, , 221–226.
Yuan,H., Quintana,J. and Dickerson,R.E. ( (1992) ) Alternative structures for alternating poly(dA-dT) tracts: the structure of the B-DNA decamer C-G-A-T-A-T-A-T-C-G. Biochemistry, , 31, , 8009–8021.
Breslauer,K.J., Frank,R., Blocker,H. and Marky,L.A. ( (1986) ) Predicting DNA duplex stability from the base sequence. Proc. Natl Acad. Sci. USA, , 83, , 3746–3750.
Leroy,J.L., Gao,X.L., Misra,V., Gueron,M. and Patel,D.J. ( (1992) ) Proton exchange in DNA–luzopeptin and DNA–echinomycin bisintercalation complexes: rates and processes of base-pair opening. Biochemistry, , 31, , 1407–1415.
Guelev,V., Lee,J., Ward,J., Sorey,S., Hoffman,D.W. and Iverson,B.L. ( (2001) ) Peptide bis-intercalator binds DNA via threading mode with sequence specific contacts in the major groove. Chem. Biol., , 8, , 415–425.
Wilson,D.W., Tanious,F.A., Fernandez-Saiz,M. and Rigi,C.T. ( (1997) ) Evaluation of drug-nucleic acid interactions by thermal melting curves. Methods Mol. Biol., , 90, , 219–240.(José Gallego*)
* To whom correspondence should be addressed at present address: Medivir UK Ltd, 100 Fulbourn Road, Cambridge CB1 9PT, UK. Email: Jose.Gallego@medivir.com
ABSTRACT
Bisnaphthalimide intercalators are anti-tumour agents composed of two planar rings linked by a flexible diazanonylene chain. The intercalated rings of three bisnaphthalimide analogues complexed to DNA are found here to undergo 180° rotating motions that do not affect the diazanonylene linker atoms bound to the major groove. These ring rotations are detected by NMR spectroscopy in a broad range of sequence contexts and duplex lengths. A comparative analysis of the frequency and activation energies of such excited states in different complexes and conditions indicates that these motions (i) are unrelated to drug dissociation; (ii) are a consequence of concerted, sequence-dependent nucleotide movements taking place on the millisecond time scale; and (iii) may occur inside the DNA duplexes. The rotation frequencies range from 2 to 25 s–1 at 25°C, depending on DNA composition and the size of the rotating rings. The detected nucleotide dynamics are likely to play an important role in the binding kinetics of the numerous proteins and drugs that require base unstacking when interacting with DNA.
INTRODUCTION
Nucleic acid dynamics influence the binding kinetics of transcription factors (1,2), endonucleases (3,4) and drugs (5–9) that intercalate or thread through the DNA double helix, and are important in DNA processing events requiring base flipping from the double helix (10,11) and strand stretching (12). Functionally relevant nucleotide motions have also been reported in ribozymes (13) and ribosome subunits (14,15). Whereas there is essential structural insight into most of these processes, a significantly smaller quantity of data is available about the direction and extent of nucleotide motions. This is especially true for time scales that are biologically (2,11,13,14,16) and pharmacologically (5–7) relevant but currently inaccessible to molecular dynamics simulations.
We recently solved the solution structure of a complex between elinafide (LU) (Figure 1a) and d(ATGCAT)2 (d6) (17). Using NMR spectroscopy, we found that the two naphthalimide rings of the drug bis-intercalate at the TpG (CpA) steps of d6, and undergo 180° rotations without affecting the 3,7-diazanonylene linker atoms bound to the DNA major groove (17) (Figure 1b). Here I have analysed these rotating motions in complexes of LU, LU-84743 (LUNH) and LU-77655 (LUNO) (18) (Figure 1a) with DNA duplexes of varying length and sequence (Figure 1c) and under different temperature and solution conditions. A comparative analysis of these complexes indicates that the rotational motion of the naphthalimide rings is detected in all sequences, and reveals sequence-dependent DNA nucleotide movements taking place on the millisecond time scale.
Figure 1. Bisnaphthalimide intercalators and DNA–bisnaphthalimide complexes. (a) Chemical structure of bisnaphthalimides. The van der Waals diameters (d) of the rotating naphthalimide rings are specified. (b) Schematic representation of a DNA–bisnaphthalimide complex and of the observed rotation of the naphthalimide rings (grey rectangles). The activated state is shown in a darker tone, and ring rotation may take place outside or inside the intercalation site. (c) The DNA–bisnaphthalimide complexes analysed in this study. The DNA sequences are self-complementary and form antiparallel duplexes containing 6 to 42 base pairs. The bisnaphthalimide molecules are shown schematically.
MATERIALS AND METHODS
Sample preparation
The DNA–bisnaphthalimide complexes were prepared as described (17) after purification of the DNA oligomers by reverse-phase high-performance liquid chromatography (HPLC) or gel electrophoresis. All samples contained 1 mM complex in 20 mM sodium phosphate buffer (pH 7), 0.2 mM EDTA, and 0 or 150 mM NaCl.
NMR spectroscopy
All NMR spectra were acquired on temperature-calibrated Bruker DRX-500 and DMX-600 spectrometers and processed with Felix 97.0 software. Each complex was analysed at several temperatures using 1H2O NOESY and a series of 2H2O NOESYs uninterruptedly collected at several mixing times (typically 0, 5, 10, 15, 20, 40, 80 and 200 ms), together with 2H2O pure exchange (19), dqf-COSY, TOCSY and ROESY experiments, all with recycle delays of 2 s. All LU, LUNH and LUNO protons and most of the aromatic, H1', H2' and H2'' DNA resonances were assigned in each of the complexes and in each of the unbound DNA duplexes studied. All sequences form the self-complementary duplexes shown in Figure 1c, although a minor hairpin form was also detected for d26.
LU ring rotation rate constants were measured at an average of nine temperatures (ranging from –6 to 56°C) in each of the eleven DNA–LU complexes and ionic strength conditions studied (Table 1; Figure 3). Depending primarily on temperature and complex size, the rates were determined from a combination of NOESY experiments, magnetization transfer data and lineshape analyses. The NOESY and magnetization transfer methods were generally applied at lower temperatures and used the H4–H9 pair of exchanging naphthalimide resonances of the LU complexes. H4 and H9 are 6.7 ? apart (Figure 1a), and the H4 resonance is shifted upfield and resolved in all complexes (Figure 2b). Magnetization transfer experiments were carried out as described before (17), at an average of two different temperatures per complex and solution condition. For NOESY measurements (on average, one per complex and solution condition), the rates were obtained using the H4 diagonal volume and the initial build-up of the H4–H9 crosspeak volume as a function of at least three mixing times, ranging from 0 to 50 ms (20). At higher temperatures, the rates were determined (at an average of six different temperatures per complex and solution condition) with the lineshape analysis program Mexico (21) using all the resolved LU naphthalimide resonances. The coupling constants of the naphthalimide system and the temperature dependence of the naphthalimide 1H chemical shifts were included in the calculations, and the reference transverse relaxation rate, T2–1, was obtained at each temperature from the linewidth of resolved aromatic DNA peaks not affected by exchange. Assuming that they are independent of temperature (Figure 3), H* and S* for LU ring rotation were obtained by fitting the rate constant (k) and temperature (T) data to the Eyring expression, ln(k/T) = ln(kb/h) + S*/R – H*/RT, where h, kb and R are Planck's constant, Boltzmann's constant and the universal gas constant, respectively. DNA–bisnaphthalimide dissociation rates were determined from NOESY experiments using the TGCA H2 resonance, shifted upfield and resolved (Figure 2b, inset) in all complexes except d22–LU, and its crosspeak to TGCA H2 in residual free DNA (Figure 2a, asterisk). All linear and non-linear (Levenberg–Marquardt) regression analyses were performed with the program Pro Fit 5.5.3. LUNH and LUNO ring rotation rates and activation parameters were determined from NOESY experiments as described for LU.
Table 1. Ring rotation rates and activation energies, dissociation rates, and thermal melting temperatures of DNA–bisnaphthalimide complexes
Figure 3. Naphthalimide ring rotation rate constants as a function of temperature and: (a) duplex length, (b) ionic strength, (c) sequence, (d) ring location and (e) ring rotating diameter. (a) d6–LU, d8–LU, d10–LU, d20–LU and d42–LU, 0 mM NaCl; (b) d10–LU and d20–LU, 0 and 150 mM NaCl; (c) d22–LU and d22a–LU, 150 mM NaCl; (d) d26–2LU, internal and external probe location, 150 mM NaCl; (e) d22a–LU, d22a–LUNH and d22a–LUNO, 150 mM NaCl. The rate constants (k) are in s–1, and the temperatures (T) in K. In (d), internal and external refer to the position of the LU rings in the d26–2LU complex (Figure 1c).
Figure 2. d20–LU 1H NMR spectra. (a) Aromatic region of the pure exchange spectrum (500 MHz, mixing time 60 ms) of d20–LU at 28°C, showing the crosspeaks between symmetrical and exchanging naphthalimide protons (Figure 1a). The exchange crosspeak identified with an asterisk connects the bound and unbound resonances of H2 of the adenine base adjacent to the naphthalimide rings in all complexes (Figure 1c). The intensity of this crosspeak is much smaller than the intensity of the crosspeaks connecting the exchanging naphthalimide ring protons, indicating that ring dissociation is slower than ring rotation. (b) 1H2O and 2H2O (inset) 1H NMR spectra (500 MHz) of d20–LU at 20 and 24°C, respectively. The LU H4 and TaHN3, TbHN3, AcH2 and GHN1 assignments of the d(A8TaGCAcTbT7)2 d20 sequence are labelled over the corresponding resonances. In (a) and (b), the axes represent 1H frequency in parts per million.
Error analysis
For NOESY and magnetization transfer experiments, the rate errors were estimated from the standard deviation of the parameters fitted by exponential or linear regression, and were within 2–6% and 5–15%, respectively. NOESY determinations included the standard deviation of the peak volumes in the calculations. For LUNH and LUNO, the average error is slightly larger (15% on average) due to smaller exchange rates. For LU lineshape analyses, the rate errors were estimated to be 10%, based on visual inspection and variation of the reference T2–1 values. The H*and S* errors (Table 1) were then obtained by propagation analysis from the standard deviations of the Eyring fits, which included the rate errors and a systematic 0.1°C temperature error in the calculations.
UV melting experiments
The thermal denaturation of the DNA–LU complexes was monitored by measuring the UV absorbance at 260 nm as a function of temperature in Beckman DU 650 or Varian Cary 500 Scan spectrophotometers. The temperature was raised to 90°C at a gradient of 0.5°C min–1 and subsequently decreased at the same rate. The concentration of DNA–LU was 0.5–1 ODU ml–1 (1–8 μM) in solutions identical to those used for NMR experiments.
RESULTS
LU ring rotation
Analysis of the NMR data confirmed that, as in d6–LU (17), the two rings of LU, LUNH and LUNO bis-intercalate into the TGCA sequence from the major groove in all complexes studied (Figure 1b). There are no indications of interaction with any other region of the oligomers. LU, LUNH and LUNO ring rotation is also observed in all complexes, as demonstrated by the presence of exchange crosspeaks between naphthalimide proton resonances (Figure 2a), detected by NOESY experiments at very short (3 ms) mixing times and confirmed by exchange experiments; the selective broadening of these naphthalimide resonances as the exchange rate increases (17); and the absence of exchange crosspeaks or broadening affecting the diazanonylene linker methylene hydrogens (17). These three observations demonstrate the occurrence of selective 180° rotations of the naphthalimide rings, which exchange between two intercalated states without affecting the linker atoms bound to the DNA major groove.
Additionally, naphthalimide ring rotation occurs at temperatures well below the melting points determined by UV and NMR thermal denaturation experiments (Table 1); the DNA base pairs are stacked and hydrogen-bonded in all complexes, as demonstrated by the observation of chemical shifts and inter-strand NOEs typical of double-helical DNA (Figure 2b); no changes in DNA or drug chemical shifts are observed at high temperatures, indicating that the complexes retain the bis-intercalated conformation near their melting points (17), and the ring rotation rates are significantly faster than the DNA–bisnaphthalimide dissociation rates (Figures 2a and 4a; Table 1).
Figure 4. (a) Comparison of d20–LU dissociation rates (blue diamonds; s–1) and d20–LU ring rotation rates (pink squares) at 150 mM NaCl and three different temperatures: 8, 16 and 24°C. Dissociation rates are less sensitive to temperature and 5- to 9-fold slower than ring rotation rates. (b) Imino proton linewidths (, Hz) as a function of temperature (T, °C) for free d10 (green lines) and d10–LU (red lines). Triangles and squares correspond to the Ta and Tb HN3 linewidths of the AAATaGCATbTT d10 sequence, respectively. The G HN1 linewidth values are connected by bare lines.
In the following sections, the frequencies and activation energies for LU, LUNH and LUNO ring rotation are analysed in different sequence, temperature and solution environments.
Duplex length
The effect of the number of flanking bases on ring rotation rates and activation energies was studied in a series of LU complexes with d6, d8, d10, d20 and d42 (Figure 1c) at 0 mM NaCl. The probe rotation rates are almost 10-fold slower in d8–LU0, with three base pairs stacked on either side of the LU rings, compared with d6–LU0, which contains only two. In contrast, the relative differences between d8–LU0, d10–LU0, d20–LU0 and d42–LU0 are smaller. In fact, the rotation rates in d42–LU0, with two full helix turns stacked on each of the LU rings, are slightly faster relative to the shorter d20–LU0 complex (Figure 3a; Table 1). The ring rotation activation enthalpies (H*) and entropies (S*) are also affected by duplex length and tend to decrease when the number of flanking base pairs increases (Table 1).
Effect of ionic strength
This was analysed by comparing d10–LU, d20–LU and d42–LU at 0 and 150 mM NaCl. Due to the screening of intermolecular electrostatic interactions, the presence of salt in the solution increases the thermal stability of the complexes, but also tends to increase the DNA–LU dissociation rates (Tm and kd, Table 1). In contrast, the rotation rates slow down significantly at 150 mM NaCl for the d10–LU system. The effect is small in the two larger complexes (Figure 3b; Table 1).
Sequence
The effect of base sequence on ring rotation was studied by analysing d22–LU and d22a–LU at 150 mM NaCl. In d22–LU the probe rings are flanked by dA10?dT10, whereas in d22a the flanking sequences are alternating, d(AT)5?d(AT)5 (Figure 1c). Homopolymeric A:T tracts adopt the relatively rigid and remarkably stable B' conformation (22), whereas alternating ones are more flexible (23) and thermodynamically less stable (24). This is reflected in the higher melting temperature of d22–LU relative to d22a–LU (Table 1). The DNA–LU dissociation rate is also slower in d22–LU than in d22a–LU (Table 1) but strikingly, the probe rotation rates are faster in the more stable d22–LU complex (Figure 3c; Table 1).
Internal versus external probe location
Since LU binds to any DNA sequence containing guanine (J. G., unpublished data), d26 was designed to also probe sequences flanked by G:C pairs (Figure 1c): two LU molecules bind to each of the TGCA sites so that, for each of them, one of the rings is in an external position, adjacent to one of the homopolymeric dA5?dT5 tracts, and the other one is in an internal position, adjacent to the alternating d(AT)6?d(AT)6 central tract. The presence of two LU molecules bound at both ends of d26 is expected to have a strong stabilizing effect, because bis-intercalating agents have been shown to clamp the nucleotides sandwiched between the drug rings and to strongly increase the lifetime of those base pairs (25). Indeed, the d26–2LU complex has the highest melting temperature (Table 1). Additionally, the different chemical environments resolved the internal and external LU H4 resonances, allowing for the comparison of the ring rotation rates at the two locations. The results are consistent with the previously observed trends: first, the internal and external rates are different, with the internally located rings rotating more slowly than the external ones (Figure 3d; Table 1); and second, the rotating motions at the internal location are favoured by a smaller H* but disfavoured by a more negative S*, whereas the opposite is true for the external rings (Table 1).
Diameter of the rotating rings
This effect was analysed by comparing d22a–LU, d22a–LUNH and d22a–LUNO at 150 mM NaCl. In contrast to LU and due to the asymmetry of their rings (Figure 1a), LUNH and LUNO give rise to two exchanging, unequally populated sets of five naphthalimide resonances (data not shown). A distinct correlation is detected between the rotation rates and the van der Waals diameter of the rotating rings (Figure 1a), with the rates slowing down markedly for LUNH and LUNO relative to the unsubstituted bisnaphthalimide (LU) (Figure 3e, Table 1).
DISCUSSION
Several conclusions can be reached from the above results. Intercalated ring rotation occurs in sequences resembling polymeric DNA and at internal locations protected from the duplex ends by base pair clamping, as can be deduced from the analysis of d6–LU, d8–LU, d10–LU, d20–LU, d42–LU and d26–2LU (Figure 3a and d; Table 1). Additionally, these rotating motions are detected at temperatures well below the UV and NMR melting points of all DNA–LU complexes (Table 1). With the possible exception of the faster rates detected in the smaller d6–LU complex, these observations rule out the possibility of ring rotation being a consequence of end effects in the DNA double helices.
A number of observations also demonstrate that intercalated ring rotation is unrelated to DNA–bisnaphthalimide dissociation. Depending on the complex and the temperature, DNA–bisnaphthalimide dissociation rates are 2- to 23-fold smaller than the ring rotation rates (Figures 2a, 4a; Table 1). Dissociation rates are also less dependent on temperature than rotation rates (Figure 4a). The d10–LU rotation rates slow down when the ionic strength is increased and the DNA duplex is stabilized (Figure 3b; Table 1, k and Tm). In contrast, the DNA–ligand dissociation rates accelerate when the ionic strength of the solution is higher (Table 1, kd), probably due to electrostatic screening of intermolecular interactions.
The observed stacked ring rotation is sequence dependent. The base sequences of d22 and d22a differ at positions that are not immediately adjacent to the naphthalimide rings, but have different conformational (22,23) and thermodynamic (24) properties. Comparison of d22–LU and d22a–LU indicates that the rotation rates reflect the sequence-dependent dynamic properties of DNA, and do not necessarily correlate with the thermal stability of the complexes or the observed DNA–ligand dissociation rates (Figure 3c; Table 1).
The rotation rate constants are affected by both the number and the sequence of flanking bases that are not adjacent to the naphthalimide rings, and depend on the internal or external location of the LU rings within the d26–2LU complex (Figure 3a, c and d; Table 1). In fact, the activation entropies (Table 1, S*) tend to become more negative as additional flanking base pairs are added or when LU ring rotation involves the internal base pairs of d26. Taken together, these observations indicate that stacked ring rotation implicates concerted base movements within the DNA–LU complexes: the probability of such concerted motions is expected to diminish with the presence of more flanking base pairs or at internal locations protected from the duplex ends by base pair clamping, hence the diminishing S* values.
Two mechanisms of intercalated ring rotation can be envisioned to explain the observations listed above. Ring rotation could take place inside the DNA duplexes, or alternatively the rings could independently slide out of the helix, flip in solution and intercalate back into DNA. The NMR experiments only provide information on the frequency and activation energies of the excited states, but not on their nature. It is therefore difficult to completely rule out one of the two possible mechanisms based on indirect evidence.
However, a number of observations suggest that ring rotation takes place inside the DNA duplexes. The external mechanism of rotation would likely involve part of the amino-alkyl linker protons, yet these protons are unaffected by naphthalimide exchange (17). Both the rates and the activation energies of LU ring rotation are exquisitely sensitive to the sequence and length of the flanking segments and to the external or internal location of the drug rings. An approximate 10-fold reduction is observed when the van der Waals diameter of the rotating rings is increased by 1 ? (Figures 1a and 3e; Table 1), and ring rotation is completely abolished in LU diimide analogues (26). A mechanism based on independent ring dissociation and rotation in the solvent could also account for the observed LU, LUNH and LUNO rate differences. However, since the three analogues share the same diazanonylene linker, one would expect a correlation between the ring dissociation/rotation frequency and the expected d22a–LU, d22a–LUNH and d22a–LUNO dissociation constants, which can be estimated from the relative Tm values of the corresponding complexes (27). This correlation is not observed (Table 1), indicating that ring rotation is unrelated to DNA dissociation.
If ring rotation takes place inside the DNA duplexes, the activated state of the rotating probes (with 9.2–10.8 ? van der Waals diameters) must require a transient interbase distance strikingly longer that the typical 3.4 ? stacking distance between bases. Stacked ring rotation may be facilitated by the spontaneous flipping (opening) of adjacent DNA base pairs in the complex. Although at significantly faster rates than the ring rotation rates reported here, base pair opening events also occur on the millisecond time scale (11,25). Ring rotation may also be assisted by spontaneous stretching and buckling nucleotide motions within the DNA duplex. Whereas the DNA base pairs are stacked and hydrogen-bonded in all complexes (Figure 2b), significant broadening of the imino proton resonances of the thymine DNA bases is observed in all sequences upon LU complex formation (Figure 4b; Supplementary Material). The observed broadening supports the occurrence of increased base motions in the bound DNA duplexes, which may facilitate in situ rotation of the intercalated rings.
The nucleotides comprising the immediate binding environment of the bisnaphthalimide ligands are the same in all complexes (TGCA), yet the ring rotation rates and activation parameters are influenced by the length, sequence and context of the adjacent nucleotide segments. Thus, irrespective of the rotation mechanism, it can be unambiguously concluded that the different naphthalimide ring rotation rates reflect the dynamic properties of each bound DNA sequence, and are facilitated by concerted nucleotide movements taking place at rates of 2–30 s–1 at 25°C. The detected dynamics are likely to play an important role in the binding kinetics of the numerous proteins and drugs that require unstacking of adjacent bases in order to intercalate or thread through the DNA double helix, and may influence DNA processing events involving separation and rejoining of complementary bases.
SUPPLEMENTARY MATERIAL
ACKNOWLEDGEMENTS
The initial stages of this work were carried out in the laboratory of Brian Reid at the Chemistry Department of the University of Washington, Seattle, USA, to whom I am grateful. I also thank Elisabeth Golden and David Loakes for help in the laboratory, Federico Gago, Miguel Bra?a and Christian Bailly for providing the bisnaphthalimide drugs, the EU Commission for a 2000–2002 Marie Curie fellowship, and the MRC for financial support.
REFERENCES
Werner,M.H., Gronenborn,A.M. and Clore,G.M. ( (1996) ) Intercalation, DNA kinking, and the control of transcription. Science, , 271, , 778–784.
Wu,J., Parkhurst,K.M., Powell,R.M., Brenowitz,M. and Parkhurst,L.J. ( (2001) ) DNA bends in TATA-binding protein–TATA complexes in solution are DNA sequence-dependent. J. Biol. Chem., , 276, , 14614–14622.
Tsutakawa,S.E., Jingami,H. and Morikawa,K. ( (1999) ) Recognition of a TG mismatch: the crystal structure of very short patch repair endonuclease in complex with a DNA duplex. Cell, , 99, , 615–623.
Horton,N.C., Dorner,L.F. and Perona,J.J. ( (2002) ) Sequence selectivity and degeneracy of a restriction endonuclease mediated by DNA intercalation. Nature Struct. Biol., , 9, , 42–47.
Fox,K.R., Brassett,C. and Waring,M.J. ( (1985) ) Kinetics of dissociation of nogalamycin from DNA: comparison with other anthracycline antibiotics. Biochim. Biophys. Acta, , 840, , 383–392.
White,R.J. and Phillips,D.R. ( (1989) ) Sequence-dependent termination of bacteriophage T7 transcription in vitro by DNA-binding drugs. Biochemistry, , 10, , 4277–4283.
Tanious,F.A., Yen,S.F. and Wilson,W.D. ( (1991) ) Kinetic and equilibrium analysis of a threading intercalation mode: DNA sequence and ion effects. Biochemistry, , 30, , 1813–1819.
Williams,H.E.L. and Searle,M.S. ( (1999) ) Structure, dynamics and hydration of the nogalamycin–d(ATGCAT)2 complex determined by NMR and molecular dynamics simulation in solution. J. Mol. Biol., , 290, , 699–716.
Onfelt,B., Lincoln,P. and Norden,B. ( (2001) ) Enantioselective DNA threading dynamics by phenazine-linked 2+ dimers. J. Am. Chem. Soc., , 123, , 3630–3637.
Roberts,R.J. and Cheng,X. ( (1998) ) Base flipping. Annu. Rev. Biochem., , 67, , 181–198.
Klimasauskas,S., Szyperski,T., Serva,S. and Wüthrich,K. ( (1998) ) Dynamic modes of the flipped-out cytosine during HhaI methyltransferase–DNA interactions in solution. EMBO J., , 17, , 317–324.
Nishinaka,T., Ito,Y., Yokoyama,S. and Shibata,T. ( (1997) ) An extended DNA structure through deoxyribose-base stacking induced by RecA protein. Proc. Natl Acad. Sci. USA, , 94, , 6623–6628.
Zhuang,X., Kim,H., Pereira,M.J., Babcock,H.P., Walter,N.G. and Chu,S. ( (2002) ) Correlating structural dynamics and function in single ribozyme molecules. Science, , 296, , 1473–1476.
Rodnina,M.V. and Wintermeyer,W. ( (2001) ) Fidelity of aminoacyl-tRNA selection on the ribosome: kinetic and structural mechanisms. Annu. Rev. Biochem., , 70, , 415–435.
Spirin,A.S. ( (2002) ) Ribosome as a molecular machine. FEBS Lett., , 514, , 2–10.
Ukiyama,E., Jancso-Radek,A., Li,B., Milos,L., Zhang,W., Phillips,N.B., Morikawa,N., King,C.Y., Chan,G., Haqq,C.M., Radek,J.T., Poulat,F., Donahoe,P.K. and Weiss,M.A. ( (2001) ) SRY and architectural gene regulation: the kinetic stability of a bent protein–DNA complex can regulate its transcriptional potency. Mol. Endocrinol., , 15, , 363–377.
Gallego,J. and Reid,B.R. ( (1999) ) Solution structure and dynamics of a complex between DNA and the antitumor bisnaphthalimide LU-79553: intercalated ring flipping on the millisecond time scale. Biochemistry, , 38, , 15104–15115.
Bailly,C., Brana,M. and Waring,M.J. ( (1996) ) Sequence-selective intercalation of antitumour bis-naphthalimides into DNA. Evidence for an approach via the major groove. Eur. J. Biochem., , 240, , 195–208.
Fejzo,J., Westler,W.M., Macura,S. and Markley,J.L. ( (1991) ) Strategies for eliminating unwanted cross-relaxation and coherence-transfer effects from two-dimensional chemical-exchange spectra. J. Magn. Reson., , 92, , 20–29.
Perrin,C.L. and Dwyer,T.J. ( (1990) ) Application of two-dimensional NMR to kinetics of chemical exchange. Chem. Rev., , 90, , 935–967.
Bain,A.D. and Duns,G.J. ( (1995) ) A new approach to the calculation of NMR lineshapes of exchanging systems. J. Magn. Reson. A, , 112, , 258–260.
Nelson,H.C.M., Finch,J.T., Luisi,B.F. and Klug,A. ( (1987) ) The structure of an oligo(dA)·oligo(dT) tract and its biological implications. Nature, , 330, , 221–226.
Yuan,H., Quintana,J. and Dickerson,R.E. ( (1992) ) Alternative structures for alternating poly(dA-dT) tracts: the structure of the B-DNA decamer C-G-A-T-A-T-A-T-C-G. Biochemistry, , 31, , 8009–8021.
Breslauer,K.J., Frank,R., Blocker,H. and Marky,L.A. ( (1986) ) Predicting DNA duplex stability from the base sequence. Proc. Natl Acad. Sci. USA, , 83, , 3746–3750.
Leroy,J.L., Gao,X.L., Misra,V., Gueron,M. and Patel,D.J. ( (1992) ) Proton exchange in DNA–luzopeptin and DNA–echinomycin bisintercalation complexes: rates and processes of base-pair opening. Biochemistry, , 31, , 1407–1415.
Guelev,V., Lee,J., Ward,J., Sorey,S., Hoffman,D.W. and Iverson,B.L. ( (2001) ) Peptide bis-intercalator binds DNA via threading mode with sequence specific contacts in the major groove. Chem. Biol., , 8, , 415–425.
Wilson,D.W., Tanious,F.A., Fernandez-Saiz,M. and Rigi,C.T. ( (1997) ) Evaluation of drug-nucleic acid interactions by thermal melting curves. Methods Mol. Biol., , 90, , 219–240.(José Gallego*)