Solution structure of a DNA duplex containing a

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letters gel. Relative velocities were calculated as extent of reaction divided by reaction time and normalized to the lowest enzyme concentration used (6.7 nM) and to the highest primer–template concentration (5 µM).

Acknowledgments This work was supported by the National Institutes of Health and by a fellowship from the Spanish Ministry of Education and Culture to J.C.M.

Juan C. Morales1 and Eric T. Kool1,2

4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

Department of Chemistry, University of Rochester, Rochester, New York 14627, USA. 2Department of Biochemistry and Biophysics, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642, USA.

1

17. 18. 19. 20.

Correspondence should be addressed to E.T.K. email: [email protected]. 21.

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Received 6 July, 1998; accepted 30 July, 1998. 1. Stryer, L. Biochemistry, 4th Ed., W. H. Freeman, New York, 89 (1995) . 2. Watson, J. D., Hopkins, N. H., Roberts, J. W., Steitz, J. A. & Weiner, A. M. Molecular biology of the gene (Benjamin/Cummings, Menlo Park; 1987). 3. Kornberg, A. & Baker, T. A., DNA Replication, 2nd ed. (W. H. Freeman, New York; 1992).

22. 23. 24. 25. 26.

Loeb, L. A. & Kunkel, T. A. Annu. Rev. Biochem. 52, 429–457 (1982). Carroll, S. S. & Benkovic, S. J. Chem. Rev. 90, 1291–1307 (1990). Echols, H. & Goodman, M. F. Annu. Rev. Biochem. 60, 477–511 (1991). El Deiry, W. S., So, A. G. & Downey, K. M. Biochemistry 27, 546–553 (1988). Joyce, C. M., Sun X. C. & Grindley N. D. J. Biol. Chem. 267, 24485–24500 (1992). Goodman, M. F. Proc. Natl. Acad. Sci. USA 94, 10493–10495 (1997). Guckian, K. M. & Kool, E. T. Angew. Chem. Int. Ed. 36, 2825–2828 (1998). Evans, T. A. & Seddon, K. R. Chem. Commun., 2023–2024 (1997). Schweitzer, B. A. & Kool, E. T. J. Am. Chem. Soc. 117, 1863–1872 (1995). Moran, S., Ren, R. X.-F. & Kool, E. T. Proc. Natl. Acad. Sci. USA 94, 10506–10511 (1997). Guckian, K. M., Morales, J. C. & Kool, E. T. J. Org. Chem., in the press. Seela, F., Bourgeois, W., Rosemeyer, H. & Wenzel, T. Helv. Chim. Acta 79, 488–498 (1996). Goodman, M. F., Creighton, S., Bloom, L. B. & Petruska, J. Crit. Rev. Bioch. Mol. Biol. 28, 83–126 (1993). Fygenson, D. K. & Goodman, M. F. J. Biol. Chem. 272, 27931–27935 (1997). Gao, J. Biophys. Chem. 51, 253–261 (1994). Kuchta, R. D., Mizrahi, V., Benkovic, P. A., Johnson, K. A. & Benkovic, S. J. Biochemistry 26, 8410–8417 (1987). Moran, S., Rex X.-F. Ren, Rumney, S. & Kool, E. T. J. Am. Chem. Soc. 119, 2056–2057 (1997). Pelletier, H., Sawaya, M. R., Kumar, A., Wilson, S. H. & Kraut, J. Science 264, 1891–1903 (1994). Eom, S. H., Wang, J. & Steitz, T. A. Nature 382, 278–281 (1996). Pelletier, H., Sawaya, M. R., Wolfle, W., Wilson, S. H. & Kraut, J. Biochemistry 35, 12742–12761 (1996). Doublié, S., Tabor, S., Long, A. M., Richardson, C. C. & Ellenberger, T. Nature 391, 251–258 (1998). Kiefer, J. R., Mao, C., Braman, J. C. & Beese, L. S. Nature 391, 304–307 (1998). Ren, R. X.-F., Chaudhuri, N. C., Paris, P. L., Rumney, S. & Kool, E. T. J. Am. Chem. Soc. 118, 7671–7678 (1996).

Solution structure of a DNA duplex containing a replicable difluorotoluene– adenine pair A nonpolar aromatic nucleoside derivative based on 2,4-difluorotoluene (F), a non-hydrogen bonding shape analog of thymidine, was recently shown to be replicated against adenine with high efficiency and fidelity. This led to the suggestion that geometric matching, potentially even in the absence of hydrogen bonding between bases in a pair, may be sufficient to direct nucleotide selection during replication. We have examined the solution structure of the F–A pair in the context of a 12 base pair DNA duplex. We find that, despite the destabilization caused by this analog, the F–A pair very closely resembles that of a T•A pair in the same context. This lends support to the importance of shape matching in replication. Electrostatic effects are widely accepted as being highly important to specific protein–DNA interactions and to the enzymatic processing of DNA. Among the more important and structurally specific of these effects are hydrogen bonds formed between protein side chains and DNA as well as between DNA bases. Many nucleoside and nucleotide analogs have been studied as probes of specific protein (enzyme)–DNA interactions1–7, and differences in binding or activity measured between natural DNA bases and non-natural analogs are often used to evaluate the importance of hydrogen bonding interactions to the activity as a whole. However, it has been pointed out that since many such analogs differ from natural nucleosides both in hydrogen bonding ability and in steric size and shape, any differences observed might be the result of electrostatic effects, steric effects or both. For this reason, nucleoside analogs that closely mimic the shape of natural nucleosides but which have greatly reduced 954

Fig. 1 a, Schematic model of a 2,4-difluorotoluene adenine pair. b, Sequence of 12-mer duplex containing a 2,4-difluorotoluene-adenine pair in the center of the consensus binding sequence14 of the SRY protein.

polarity have been examined8,9. One of these, a difluorotoluene nucleoside (F) that mimics thymidine (T), has been studied recently as a probe of DNA replication mechanisms10,11. Moran et al.10 have reported studies in which F is tested as a template for replication by the Klenow fragment of E. coli DNA polymerase I. It was shown that dATP is inserted efficiently opposite this analog, while dCTP, dTTP and dGTP are not. Conversely, it was subsequently shown that the nucleoside triphosphate derivative of F (dFTP) is selectively inserted opposite A in a template strand11. These results were surprising, since base–base hydrogen bonding has long been considered to be a primary factor governing the insertion of nucleotides by DNA polymerases, and yet analog F is a nonpolar molecule that forms very weak hydrogen bonds12. Indeed, F has been found to destabilize duplexes in which it is paired with adenine11. nature structural biology • volume 5 number 11 • november 1998

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letters c

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a

Fig. 2 a, Thermal denaturation curves of F6–A19 duplex (red) and a control duplex (blue) containing thymine in place of 2,4-difluorotoluene. The buffer contains 1 M Na+, 10 mM phosphate (pH 7.0), 0.1 mM EDTA, and 37 µM of duplex. b, Imino proton spectra of the F6–A19 duplex at 9, 18, and 27 oC. The imino protons of the terminal base pairs are visible at 3 oC, but disappear by 9 oC due to exchange with water. c, Base to H1' region of a NOESY spectrum at 375 ms for the F6–A19 duplex at 20 oC.

An alternative explanation for these observations is a steric selection model, in which base pair shape complementarity serves as a chief factor influencing the selection of nucleotides for incorporation during replication. Although studies have shown that the free nucleoside F acts as a close structural and conformational mimic of thymidine13, in light of steric versus hydrogen bonding models it is important to evaluate whether an F–A pair, in the context of duplex DNA, resembles a T•A base pair in terms of local and extended structure effects. Based on the instability of the F–A pair, it would be reasonable to anticipate significant disruption of duplex structure by this substitution. We have investigated this question by studying an F–A pair in the context of a 12 base pair DNA duplex (Fig. 1) in solution by two-dimensional NMR combined with restrained molecular dynamics. The results are important in determining whether F–A can accurately mimic the T•A in geometry, a question that is central to evaluating relative influences of steric and hydrogen bonding effects in replication. In addition, the findings suggest that nucleoside F may serve as a useful probe of structure and hydrogen bonding in varied DNA and protein–DNA complexes. nature structural biology • volume 5 number 11 • november 1998

Unstable, nonselective pairing by the F analog A 12-mer duplex containing a central F–A pair (Fig. 1) was constructed; the sequence corresponds to the consensus binding site of the transcription factor SRY14. To evaluate the inherent pairing preferences of F in this sequence context we also prepared three duplexes containing F–G, F–T, and F–C mismatches at the same position. Four duplexes containing a natural T•A pair or T–G, T–T, T–C mismatches were made for comparison. Thermal denaturation studies of these sequences in a pH 7.0 buffer (100 mM NaCl, 10 mM phosphate, 0.1 mM EDTA, 3 µM DNA) gave Tm values of 36 oC for the F–A duplex and 35, 34 and 31 oC for the F–G, F–T and F–C sequences respectively (data not shown). The corresponding free energies (37 oC) were -8.1, -7.7, -7.3 and -7.1 kcal mol–1 for the same four cases. Thus, the results show that there is little if any inherent pairing preference of F for A. The denaturation data for the natural pair and mismatches in this context were significantly different: the Tm (free energy) values were 50 oC (-11.5 kcal mol–1) for the T•A duplex and 43 oC (-9.5 kcal mol–1), 36 oC (-8.1 kcal mol–1), and 33 oC (-7.3 kcal mol–1) for the T–G, T–T, and T–C mismatched duplexes respectively, showing a strong preference for pairing of T with A. Thus, the F–A pair is nonse955

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letters Fig. 3 a, Superposition of 12 randomly selected final structures, six of which started from B-form DNA and six from A-form DNA. b, Stereoviews of the central base pairs of the F6–A19 duplex. Top, view from the major groove, showing the F6(yellow)–A19(green) pair and its immediate neighbors (blue). Bottom, cutaway view along the helical axis. Figure produced using the MidasPlus package31,32 from the Computer Graphics Laboratory, University of California, San Francisco.

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lectively formed and is as destabilizing as natural base mismatches. This degree of destabilization is consistent with reported values for other duplexes containing F–A pairs11. At higher DNA concentration in a 1 M NaCl buffer, the F–A duplex was found to have a Tm of 51 oC while a control duplex with T opposite A had a Tm of 62 oC (Fig. 2a).

a

b

The F–A duplex adopts a B-form conformation The chemical shifts of the imino protons of the F6–A19 duplex (Fig. 2b) indicate the formation of Watson-Crick base pairs for all A•T and G•C partners. The chemical shift of the T5 imino proton that is adjacent to the F–A pair in the modified duplex is 0.35 p.p.m. upfield to the equivalent proton in the control duplex. All other imino protons in the F6–A19 duplex are within 0.1 p.p.m. of the corresponding imino protons in the control duplex. The T5(H3) proton broadens at a lower temperature than in other T•A base pairs (Fig. 2b). The broadening of the T5 imino proton is consistent with a decrease in the lifetime of base-pair opening for the T5•A20 base pair, although catalyzed exchange experiments are required to quantify lifetimes15,16. The transient opening of the F6–A19 pair is of interest, but the lack of an imino proton for this pair precludes determination of lifetimes by standard techniques. Uninterrupted NOE connectivities are observed in the base to H1' (Fig. 2c), base to H2'/H2", and base to H3' regions of the NOESY spectra for the F6–A19 duplex as well as the control duplex (data not shown), allowing proton assignments to be made. A comparison of NOE intensities in these regions shows that all bases adopt an anti conformation. The anti conformation of the F6 base within the duplex is consistent with X-ray and solution studies in which the F nucleoside adopts an anti conformation13. Comparison of the H1' to H2'/H2" region of a DQF-COSY spectrum of the F6–A19 duplex with that of the control duplex shows similar crosspeak patterns for all resolvable resonances, including the C18–A20 and the T5–G7 sugars, confirming similar sugar puckers throughout both duplexes. Furthermore, all resolvable cross peaks in this region show a separation of greater than 14 Hz between the outer two peaks along the H1' frequency axis, which indicates an S sugar pucker17–19. The nonexchangeable proton chemical shifts of the F6–A19 duplex and the control were compared (data available from the author) and found to be similar with the exception of the F6 nucleoside and its immediate neighbors. The similarity of the chemical shifts and DQF-COSY crosspeak patterns is consistent with both duplexes adopting similar conformations. The 31P resonances for the F6–A19 duplex lie within a 1 p.p.m. range, which is also consistent with a B-form duplex20. 956

F6 and A19 stack within the helix In the NOESY spectrum in D2O at 20 oC the F6(H3) proton shows NOEs to the A19(H2) proton, as well as to the A20(H2) proton in the adjacent pair (Fig. 2c). The F6(Me) protons show NOEs to both the T5(Me) and T5(H6) protons, while the T5(Me) protons show NOEs to both the F6(Me) and F6(H6) protons. In a NOESY WATERGATE experiment at 3 oC, the F6(H3) proton exhibits crosspeaks both to the T5 and G7 imino protons of the flanking base pairs. Taken together, these data show that the predominant conformation has the F6 base stacked within the duplex across from A19. Heteronuclear 19F NOE experiments confirm the location of the 2,4 difluorotoluene base analog. The 19F spectrum of the F6–A19 duplex at 25 oC shows two major peaks (data not shown). Saturation of the F6(19F4) fluorine resonance produced small NOEs both to the A19(H2) and A20(H2) protons as well as a large NOE to the F6(Me) protons, allowing the fluorines to be assigned. Upon saturation of the F6(19F2) fluorine resonance, qualitatively large NOEs were seen both to A19(H2) and A20(H2) protons. In addition, large NOEs were also seen from the F6(19F2) fluorine resonance to the F6(H1'), F6(H2'), and F6(H2") protons, consistent with an anti conformation. In H2O solution both the G7 and T5 imino protons exhibited NOEs when either the F6(19F2) or the F6(19F4) fluorine resonature structural biology • volume 5 number 11 • november 1998

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nances were irradiated. These NOEs also are consistent with a Table 1 Structural statistics for the difluorotoluene containing predominant conformation in which the 2,4 difluorotoluene duplex base analog is stacked in the duplex. Structural restraints Structural calculation and structural features NMR data sets were collected and analyzed to obtain 252 NOE distance restraints and 136 dihedral angle restraints for the 12mer duplex. Nonexchangeable proton distance restraints were obtained by inputting volumes from NOESY data at several mixing times into MARDIGRAS21–23. The MARDIGRAS output contains upper and lower bounds that were used with hydrogen bonding and dihedral angle restraints (Table 1) as input for restrained molecular dynamics calculations. Both Aform and B-form 12-mer duplexes containing F6–A19 base pairs were used as initial structures. The convergence of the initial structures is illustrated in Fig. 3a, and the structural statistics are given in Table 1 and the Methods. The final coordinates from 50 molecular dynamics simulations were averaged and the resulting structure was energy minimized to give the structure shown in Fig. 3b. In all the calculated structures the F6 base is stacked within the duplex opposite the A19 base. The structures found at the completion of molecular dynamics, energy minimization (MD/EM) computations exhibit a small change in the alignment of the A19 and F6 bases when compared to the starting structure. Initially, the A19 and F6 bases were arranged such that the A19(N1):F6(H3):F6(C3) atoms were colinear (at an angle of 180o) as found in a canonical A•T base pair. In the final structures the A19(N1) : F6(H3) : F6(C3) angle has an average value of 140o (Fig. 3b), reflecting a small shift of the F6(H3) proton from a colinear to a staggered arrangement. Both colinear and staggered conformations are consistent with NMR restraints. The chemical shift of the F6(H3) proton of the 2,4 difluorotoluene base moves 0.30 p.p.m. downfield as the temperature is lowered from 30 to 3 oC, which contrasts with the A19(H2) proton that moves upfield slightly (+0.04 p.p.m.) over this temperature range. The chemical shift of the F6(H3) proton in the single strand containing the 2,4 difluorotoluene base moves upfield (+0.07 p.p.m.) as the temperature is lowered (data not shown). Thus the downfield movement of the F6(H3) proton in the F6–A19 duplex as the temperature is lowered from 30 oC to 3 oC reflects changes in the local environment of the F6(H3) proton within the duplex. The downfield movement in the chemical shift of the F6(H3) proton may be due to a subtle change in the stacking of adjacent base pairs, or to the close proximity of the F6(H3) proton to the A19(N1) nitrogen. Discussion The difluorotoluene nucleoside analog is of interest because it is efficiently and selectively replicated despite its apparently very poor ability to form hydrogen bonds. Calculations have predicted that this pair forms only weak hydrogen bonds in the gas phase24, and experiments in the context of duplex DNA show that pairing between these structures is too weak to be measurable in water11. The present denaturation data confirm this, since there is no selectivity of pairing of F with A over the other bases. In addition, the pairing of F and A in a DNA duplex is destabilizing despite the fact that it causes little distortion to the structure. This is not due to poor stacking of F, since separate studies have shown that it, in fact, stacks more strongly than T or A25. Thus, we attribute this destabilization to the high cost of desolvation of the hydrogen bonding groups nature structural biology • volume 5 number 11 • november 1998

Distance restraints Total Intraresidue Interresidue Exchangeable Nonexchangeable Hydrogen-bonding Dihedral angle restraints Violations of experimental restraints in the final structure 10% restraint file1 A-form DNA was used as a starting structure Distance violations (>0.1 Å) Total Intraresidue Interresidue Dihedral violations(> 2o) B-form DNA was used as a starting structure Distance violations(>0.1 Å) Total Intraresidue Interresidue Dihedral violations(>2o) 15% restraint file1 B-form DNA was used as a starting structure Distance violations(>0.1 Å) Total Intraresidue Interresidue Dihedral violations(>2o) Atomic r.m.s. differences4 (Å) A-form 10% and B-form 10% 0.80 B-form 10% and B-form starting structure 2.34

314 112 202 25 227 62 136

2 43 1 3 1 2 43 0 4 1

2 23 2 0 1

Restraint file was generated with at least ±X% of the average distance as upper and lower bounds. 2Energy minimized average structure from 50 calculations. 3No violations >0.2 Å. 4Average pairwise r.m.s differences were calculated using energy minimized average structures generated from 50 calculations. 1

of adenine when juxtaposed with difluorotoluene. Also consistent with this are results of octanol:water partitioning studies with nucleosides dA and dF, which show that dA is hydrophilic (logP = -0.89), while dF is quite lipophilic (logP = 1.39) (unpublished results). The difluorotoluene–adenine pair has been shown to be replicated with high efficiency and fidelity by the Klenow fragment of DNA polymerase I10,11. This observation has led us to propose that shape selection may be a chief factor governing nucleotide insertion by DNA polymerase10,11, even in the absence of hydrogen bonds in a given pair. The structural data make it clear that the F–A pair can adopt a geometry closely approximating that of Watson-Crick base pairs, which is consistent with the idea that the enzyme may process the F–A pair correctly in large part because the ‘bases’ have complementary shapes in the context of the B-form duplex. The observed destabilization caused by pairing of F with natural bases may have led one to expect significant deviation from B-form structure, including the possibilities of a syn glycosidic orientation, extrahelical location of F or its partner, or distortion of neighboring pairs. Despite this, the duplex in 957

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letters Fig. 3b is B-form in nature and shows the F6 base stacked within the helix opposite the A19 base in standard anti/anti configuration. Comparison of the F–A duplex NMR data with that of the T•A duplex data suggests the formation of similar structures. The fact that the F analog is stacked in the helix despite the destabilization may be due to its strong stacking tendency, which has been demonstrated by substitution at a dangling position in short DNA duplexes25. The fact that the difluorotoluene–adenine pair closely resembles a thymine–adenine pair in this context lends confidence that the difluorotoluene ‘base’ can act as a probe of the importance of Watson-Crick hydrogen bonding in the absence of significant steric effects. This has not been possible with other known hydrogen-bonding-impaired nucleoside analogs that in most cases differ substantially in steric size and shape as well in hydrogen bonding ability. Thus, nucleoside F is likely to continue to be useful in probing polymerases, and the F–A pair may serve as a useful nonpolar T•A pair surrogate in testing major and minor groove interactions by DNA binding proteins as well. In addition to its utility as a probe of electrostatic interactions, the difluorotoluene nucleoside may also have significant use as a structural probe. C-5 fluorinated pyrimidines have provided useful information on structural interactions in the major groove26. The difluorotoluene nucleoside contains 19F nuclei in both the major and minor grooves and therefore expands the utility of fluorinated base analogs in probing macromolecular interactions. Methods Sample preparation. The 2,4 difluorotoluene phosphoramidite was synthesized27 and incorporated into a DNA 12-mer using an Applied Biosystems 392 DNA synthesizer. The unmodified and modified oligonucleotides were synthesized in trityl-on mode and purified by HPLC28. The purity was checked by PAGE and found to be greater than 95%. The strands were mixed while monitoring the integrated intensity of the base protons in D2O at 65 oC to obtain a 1:1 ratio. The sample was dissolved in buffer (100 mM NaCl, 1 mM EDTA, and 10 mM sodium phosphate, pH 7.4) at a duplex concentration of 3 mM. Thermal denaturation. The absorbance at 260 nm was monitored as a function of temperature (0–90 oC) on a Gilford 250 spectrophotometer. The duplexes were dissolved in 100 mM or 1 M NaCl, 10 mM sodium phosphate, and 0.1 mM EDTA at a pH of 7.00 to give a final duplex concentration of 3 µM or 38 µM. The curve was fit assuming a two state model29,30. NMR spectra. Experiments were performed on a Varian 500 MHz NMR spectrometer. Proton assignments were made using standard two-dimensional techniques, including NOESY, DQF-COSY, and NOESY WATERGATE. Data processing was done using Felix 95 (BIOSYM/Molecular Simulations). Proton assignments for both the difluorotoluene modified duplex as well as the unmodified duplex are available as supplementary material that can be requested from the corresponding author. All structural restraints involving nonexchangeable protons were derived from NOESY data acquired at 20 oC with mixing times of 75, 150, 225, 300 and 375 ms. Restraints for exchangeable protons were derived from NOESY WATERGATE data acquired at 3 oC with a mixing time of 150 ms. Backbone conformation and sugar pucker were investigated at 20 oC using 1D 31P and DQF-COSY experiments respectively. One dimensional 1H{19F} heteronuclear NOE experiments were performed using a Varian triple resonance probe (N15, C13 and H1). Experiments were performed in D2O at 25 oC using the S2PUL pulse sequence, and in H2O solution the BINOM pulse sequence was used with 1:1 water suppression at 3 oC. Because long irradiation times were employed in these experiments the NOEs observed were not incorporated into the restraint file.

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Restraint generation. The 227 restraints involving nonexchangeable protons were derived using MARDIGRAS21–23. From each experimental volume a distance was derived using an iterative relaxation matrix for each mixing time. The distances were averaged and the standard deviations were used to calculate the upper and lower bounds. A minimum of 10% was used for the standard deviation. A second set of distance restraints was generated using a minimum standard deviation of 15%. Twenty-five exchangeable proton restraints were given an upper bound of 5 Å and a lower bound of 1.5 Å. Dihedral angle restraints were used to preserve a right-handed DNA helix. The δ torsion angle was restrained to 110° ≤ δ ≤ 170o (C2'-endo sugar pucker) based on a greater than 14 Hz coupling for the outer two peaks in the 1' frequency dimension of a DQF-COSY spectrum17–19. A total of 252 NOE distance restraints and 136 dihedral angle restraints were used with 50 kcal mol–1 Å–2 energy terms when applied full scale. In addition, 62 restraints were included to maintain Watson-Crick pairing for base pairs 1–4 and 8–1223, consistent with NOE data. Molecular dynamics. Starting structure generation and visualization of calculated structures were performed using InsightII 95.0.6 (BIOSYM/Molecular Simulations). All structural calculations and analyses were performed using the Discover, Analysis and NMR Refine modules of InsightII. The AMBER forcefield in InsightII was modified to accommodate the fluorine atom using parameters from the AMBER95 forcefield in MACROMODEL v. 6.0 (C. Still, Columbia University). The partial charges for the 2,4 difluorotoluene nucleoside were assigned by ab initio methods at the MP2/6-31G(d) level using Gaussian 94 (Frisch, M.J. et al., Gaussian Inc.). The partial charges used in the molecular dynamics simulation for the difluorotoluene monomer are the following; C1 = -0.15, C2 = 0.52, F2 = -0.38, C3 = -0.37, H3 = 0.25, C4 = 0.50, F4 = -0.38, C5 = -0.18, Me = -0.65, MeH’s = 0.23, C6 = -0.18, H6 = 0.24, C1' = 0.13, C1'H = 0.21. Molecular dynamics simulations consisted of an initial 5,000 fs of equilibration followed by 10,000 fs of dynamics starting with random velocities at 1,000 K. The force constants were scaled from an initial scaling factor of 0.1 to a final factor of 1. The system was then equilibrated at 300 K for 5,000 fs and subjected to 10,000 fs of dynamics with a restraint scaling factor of 1. The third phase consisted of 5,000 iterations of steepest descents energy minimization at 300 K with a restraint scaling factor of 1. Lastly, conjugate gradient energy minimization was run with a distant-dependent dielectric of 1 (the electrostatic terms were turned off in all previous phases) and a restraint scaling factor of 1 for 5,000 iterations at 300 K. All phases of the calculation were run with a Lennard-Jones potential with a cutoff of 20 Å. The global covalent scale and global nonbonded scale were set to 1 in each phase. 50 calculations were performed using B-form DNA as the starting structure while using the 10% restraint file. An additional 50 calculations were performed with a B-form starting structure using the 15% restraint file. A final set of 50 calculations were performed with an A-form starting structure and the 10% restraint file. For each of the three sets of calculations all 50 structures were averaged and subjected to 100 iterations of steepest descents energy minimization with full restraints to generate the final structure. Coordinates. The atomic coordinates have been deposited in the Brookhaven Protein Data bank (accession number 1bw7)

Acknowledgements We thank the NIH for support. We thank M. Fountain for many helpful discussions.

Kevin M. Guckian, Thomas R. Krugh and Eric T. Kool Department of Chemistry, University of Rochester, Rochester, New York 14627, USA. Correspondence should be addressed to T.R.K. e-mail: [email protected] or E.T.K. e-mail: [email protected] Received 24 April, 1998; accepted 24 August, 1998.

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letters 1. Lesser, D.R., Kurpiewski, M.R. & Jen-Jacobson, L. Science 250, 776–786 (1990). 2. Brennan, C.A., Van Cleeve, M.D. & Gumport, R.I. J. Biol. Chem. 261, 7270–7278 (1986). 3. Seela, F. & Driller, H. Nucleic Acids Res. 14, 2319–2332 (1986). 4. Ono, A., Sato, M., Ohtani, Y. & Ueda, T. Nucleic Acids Res. 12, 8939–8949 (1984). 5. Newman, P.C. et al. Biochemistry 29, 9891–9901 (1990). 6. Aiken, C.R. & Gumport, R.I. Meth. Enz. 208, 433–457 (1991). 7. Smith, S.A., Rajur, S.B. & McLaughlin, L.W. Nature Struct. Biol. 1, 18–22 (1994). 8. Schweitzer, B.A. & Kool, E.T. J. Org. Chem. 59, 7238–7242 (1994). 9. Chaudhuri, N.C., Ren, X.F.R. & Kool, E.T. SYNLETT, 341–347 (1996). 10. Moran, S., Ren, X.F.R. & Kool, E.T. J. Am. Chem. Soc. 119, 2056–2057 (1997). 11. Moran, S., Ren, X.F.R. & Kool, E.T. Proc. Natl. Acad. Sci. USA 94, 10506–10511 (1997). 12. Schweitzer, B.A. & Kool, E.T. J. Am. Chem. Soc. 117, 1863–1872 (1994). 13. Guckian, K.M. & Kool, E.T. Angew. Chem. Int. Ed. 36, 2825–2828 (1998). 14. Harley, V.R., Lovell-Badge, R. & Goodfellow, P.N. Nucleic Acids Res. 22, 1500–1501 (1994). 15. Leroy, J.L., Kochoyan, M., Huynhdinh, T. & Gueron, M. J. Mol. Biol. 200, 223–238 (1988).

16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29.

A novel DNA-binding motif shares structural homology to DNA replication and repair nucleases and polymerases

dead ringer (dri) which was found to be developmentally regulated and is essential for the development of Drosophila embryos5. A homologous DNA-binding domain also exists in SWI1, a factor directly involved in DNA binding in the SWI/SNF complex6. This complex interacts with chromatin and alters chromatin structure during gene transcription7,8. This DNAbinding sequence motif also exists in RBP1 and RBP2, SMCX, SMCY, jumonji and Caenorhabditis elegans cosmid T23D89-13. The solution structure of the DNA-binding domain of Mrf-2 (residues 14–120 of Mrf-2 with full length molecular weight of 106,000 Mr) was determined and is shown in Fig. 2. Statistics of the NMR structures are shown in Table 1. The structure of the Mrf-2 domain consists of helices and loops. No β-strands have been observed. There are six helices encompassing residues 14–29, 48–56, 60–66, 69–75, 85–94, 100–105, corresponding to H1–H6, respectively (Figs 1,2a). The helical regions of the Mrf-2 domain have been well defined. H1 interacts with H6, H2 and H4, connected by H3, have a helical crossing angle of ~130 o. H4 and H5 are connected by a 10-residue disordered loop with a helical crossing angle of ~150o. Residues 95–100, which connect H5 and H6, form a distorted helical turn that causes a kink between H5 and H6. Based on characteristic dαN(i, i+2), dαN(i, i+3) and dαN(i, i+4) NOEs14, H6 forms a 310-helix, whereas H1, H2, H4 and H5 are α-helices. H3 is the most flexible helix with no detectable amide protons after exchanging with D2O for three days. In the 310-helix, NOE constraints are also more consistant with backbone Oi–NHi+3 hydrogen bonding. Five helices, H1, H2, H4, H5 and H6, have stable hydrogen bonds indicated by slowly-exchanging amide protons at these positions (Fig. 1). The loop regions and the C-terminus are flexible in solution. Backbone 15N-1H NOE measurements (Fig. 2c) show that these regions have smaller or negative values of NOEs indicative of conformational flexibility. Backbone 15N R2 (1/T2) measurements (Fig. 2c) show larger values for residues in the loop encompassing residues 30–47, suggesting conformational exchange in this region. Conformational flexibility in these regions is also shown by averaged 3JHNα coupling constants and fast exchange of backbone amide protons. The three-dimensional structure is stabilized by a hydrophobic core. The residues in the hydrophobic core are Leu 20, Ile 23, Tyr 24, Met 27, Thr 32, Ile 34, Met 50, Ala 53, Ala 54, Leu 57, Ile 63, Trp 69, Ile 72, Leu 76, Ala 85, Ala 86, Thr 89, Tyr 93, Ile 97 and Leu 98. Most of these residues are conserved and occur, in many cases, in helical regions; at every third and/or fourth position in the primary sequence (Fig. 1). A few Gly and Pro residues, such as Gly 58, Gly 59 and Pro 80 are also conserved.

A novel class of DNA-binding domains has been established from at least sixteen recently identified DNA-binding proteins. The three-dimensional structure of one of these domains, Mrf-2, has been solved using NMR methods. This structure is significantly different from known DNA-binding domain structures. The mechanism of DNA recognition by this motif has been suggested based on conserved residues, surface electrostatic potentials and chemical shift changes. This new DNA-binding motif shares structural homology with T4 RNase H, E. coli endonuclease III and Bacillus subtilis DNA polymerase I. The structural homology suggests a mechanism for substrate recognition by these enzymes. Protein–DNA interactions play central roles in many cellular processes. While many DNA-binding proteins, including transcription factors, bind to specific DNA sequences, other proteins exhibit no apparent sequence specificity but instead bind to unique DNA structures. This latter group of proteins includes enzymes involved in DNA replication and repair processes, enzymes that change the topology of DNA, histones, resolvases and integrases. The enormous number of DNA binding proteins isolated and characterized to date bind to DNA through only a small number of structurally independent motifs, such as the helix-turn-helix, zinc-finger, leucine zipper, helix-loop-helix, and β-ribbon DNA binding domains1,2. A given motif may bind to a specific DNA sequence in one protein and non-specifically in another protein. A new DNA-binding motif that does not resemble any previously identified motifs has been defined by 16 different proteins, whose DNA-binding domain sequences are aligned in Fig. 1. Mrf-1 and Mrf-2 are transcription factors which recognize the modulator region of the major immediate-early gene of human cytomegalovirus (HCMV)3. This DNA-binding sequence motif was first described in the mouse protein bright which is necessary for maturation-induced µ-heavy chain transcription in B-cells4. This motif has also been characterized in the Drosophila protein nature structural biology • volume 5 number 11 • november 1998

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