sequence specific dna binding of ets-1 transcription factor

Sequence Specific DNA Binding of Ets-1 TranscriptionFactor: Molecular Dynamics Study on the EtsDomain–DNA Complexes

Satoshi Obika, Swarnalatha Y. Reddy and Thomas C. Bruice*

Department of Chemistry andBiochemistry, University ofCalifornia, Santa Barbara, CA93106, USA

Molecular dynamics (MD) simulations for Ets-1 ETS domain–DNA com-plexes were performed to investigate the mechanism of sequence-specificrecognition of the GGAA DNA core by the ETS domain. Employing thecrystal structure of the Ets-1 ETS domain–DNA complex as a startingstructure we carried out MD simulations of: (i) the complex between Ets-1 ETS domain and a 14 base-pair DNA containing GGAA core sequence(ETS–GGAA); (ii) the complex between the ETS domain and a DNA hav-ing single base-pair mutation, GGAG sequence (ETS–GGAG); and (iii) the14 base-pair DNA alone (GGAA). Comparative analyses of the MD struc-tures of ETS–GGAA and ETS–GGAG reveal that the DNA bendingangles and the ETS domain–DNA phosphate interactions are similar inthese complexes. These results support that the GGAA core sequence isdistinguished from the mutated GGAG sequence by a direct readoutmechanism in the Ets-1 ETS domain–DNA complex. Further analyses ofthe direct contacts in the interface between the helix-3 region of Ets-1 andthe major groove of the core DNA sequence clearly show that the highlyconserved arginine residues, Arg391 and Arg394, play a critical role inbinding to the GGAA core sequence. These arginine residues make biden-tate contacts with the nucleobases of GG dinucleotides in GGAA coresequence. In ETS–GGAA, the hydroxyl group of Tyr395 is hydrogenbonded to N7 nitrogen of A3 (the third adenosine in the GGAA core),while the hydroxyl group makes a contact with N4 nitrogen of C40 (thecomplementary nucleotide of the fourth guanosine G4 in the GGAGsequence) in the ETS–GGAG complex. We have found that this differencein behavior of Tyr395 results in the relatively large motion of helix-3 in theETS–GGAG complex, causing the collapse of bidentate contacts betweenArg391/Arg394 and the GG dinucleotides in the GGAG sequence.

q 2003 Elsevier Ltd. All rights reserved.

Keywords: molecular dynamics; Ets-1; ETS domain; transcription factor;protein–DNA complex*Corresponding author

Introduction

The ETS protein family contains more than 45eukaryotic transcription activators and inhibitors,such as Ets-1, PU.1, Fli-1, GABPa, SAP-1, TEL andElk-1.1 – 3 Members of this family play an importantrole in normal cell proliferation and differentiation.The DNA rearrangement and/or overexpression ofets gene have been known to lead totumorigenesis.4 In order to regulate geneexpression, the ETS family of proteins bind to a

consensus DNA sequence centered on the coresequence 50-GGA(A/T)-30 through the highly con-served DNA-binding domain.3 The DNA-bindingdomain for ETS proteins, termed ETS domain, isabout 85 amino acid residues in length and formsa winged helix-turn-helix motif consisting of threea-helices and four b-strands. The recent X-ray5 – 10

and NMR11 – 13 studies of the ETS domain–DNAcomplexes have shown that the helix-3 in thewinged helix-turn-helix motif binds in the majorgroove of the consensus DNA sequence.

In the crystal structure of Ets-1 ETS domain–DNA [d(TAGTGCCGGAAATGT)2] complex (PDBcode: 1K79), two arginine residues, Arg391 andArg394, which are in the helix-3 region and

0022-2836/$ - see front matter q 2003 Elsevier Ltd. All rights reserved.

E-mail address of the corresponding author:[email protected]

Abbreviation used: MD, molecular dynamics.

doi:10.1016/S0022-2836(03)00726-5 J. Mol. Biol. (2003) 331, 345–359

conserved among the ETS family, make bidentateinteractions with G1 and G2, respectively (Figure1).5 However, the pattern of these hydrogen bondsis not maintained in the crystal structures of otherETS domain–DNA complexes.7,9,10 In addition, theinteraction between the arginine residues, Arg391and Arg394, and the consensus DNA sequence isnot observed in the NMR study on Ets-1 ETSdomain–DNA complex.12,13

On the other hand, a few studies have proposeddirect contacts of the amino acid residues in ETSdomain with the AA region (þ3 and þ4 positions)in the GGAA core. For example, an X-ray study ofthe Ets-1 ETS domain–DNA complex indicatedwhat would be a vital role of hydrophobic inter-action between the phenyl ring of Tyr395 and 5-methyl group of T40 or T50.

5 However, this type ofinteraction was not observed in other ETSdomain–DNA complexes,6 nor is the tyrosine resi-due conserved in other ETS family proteins suchas PU.1 and TEL.9,10 Thus, the precise molecularmechanism that clearly explains the sequence-specific GGAA recognition by ETS domain is stilllacking.

The phosphate groups of DNA adjacent to thecore sequence GGAA have contacts to the wingedsegment and the turn region between helix-2 andhelix-3 of the ETS domain. It was reported that theneutralization of anionic phosphate charges onone face of DNA resulted in the DNA bending,probably due to the electrostatic repulsions of theremaining anionic charges.14 – 16 In fact, DNA bend-ing was observed in the crystal structures of theETS domain–DNA complexes.5 – 10 It was also sup-posed that the conformational change of DNAcaused by the DNA bending would serve to pro-vide effective GGAA core recognition by the helix-3 of the ETS domain. However, the bending anglesof DNA previously reported in the X-ray crystallo-graphic analyses of ETS domain–DNA complexesvary from one system to another.5 – 10 Thus,

additional investigation is required in order to clar-ify a common role of the DNA bending in thesequence-specific binding of the ETS domain.

Significant developments have been made in thelast few years in the procedures of moleculardynamics (MD). Improvements in AMBER,17

CHARMM18,19 and GROMOS20 force fields andeffective treatment of long-range electrostatic inter-actions by using particle mesh Ewald (PME)method,21 explicit inclusion of solvent and ionshave opened the possibilities for accurate determi-nation of protein and DNA structures.22 Besidesavailability of supercomputers have enabled toundertake simulations on a nanosecond (ns) time-scale which expanded conformational samplingand eventually to elucidate biomolecular inter-actions. Further understanding of the molecularmechanism of sequence-specific DNA binding ofthe ETS domain is likely to provide novel clues forthe design of drugs that bind to inhibit the inter-action between the ETS domain and DNA. Wereport here 3.5–3.9 ns MD simulations of two Ets-1 ETS domain–DNA complex systems. The aminoacid sequence of Ets-1 ETS domain and the DNAsequences used in this study are shown in Figure 2.The binding activity of the ETS domain is knownto be higher than that of the whole ETS-1protein.23 – 27 Therefore, the ETS domain–DNAcomplex would be a good model system for MDsimulation. The first system has the ETS domain(103 amino acid residues) of Ets-1 protein (Figure2(a)) and the high affinity 14 base-pair DNA con-taining GGAA core (þ1 to þ4, Figure 2(b))sequence, while the second one has a low affinityDNA involving a mutation of a single base-pair,GGAG sequence. In the text, we refer to these com-plexes as ETS–GGAA and ETS–GGAG, respect-ively. In addition, results from the MD simulationof the 14 base-pair DNA having the GGAA coresequence (referred as GGAA) are also discussedfor comparison.

Figure 1. Crystal structure of Ets-1 ETS domain–DNA complex (PDB code: 1K79): (a) the whole structure, (b) theclose-up view of the ETS domain–DNA nucleobase contact site (helix-3 and GGAA core sequence).

346 MD Simulations of Ets-1 ETS Domain–DNA Complexes

Results

The root-mean-square deviations (RMSD) of theprotein backbone and DNA heavy-atoms withrespect to the minimized structures of ETSdomain–DNA complexes and DNA (GGAA) aregiven in Figure 3. During the simulation, theRMSD values of the protein fluctuated around1.16–2.12 A in ETS–GGAA and 0.92–1.55 A inETS–GGAG (Figure 3(a)), while those of DNA inETS–GGAA and ETS–GGAG are from 1.06 A to2.18 A and from 1.12 A to 1.96 A, respectively(Figure 3(b)). The plots indicate the stability atabout 900 ps, except in the ETS–GGAA proteinstructure. The DNA structure of GGAA exhibitsrelatively large RMSD values, compared to theETS–GGAA and ETS–GGAG complexes (greenline in Figure 3(b)). This indicates the absence ofETS domain would cause conformational changesin the DNA structure.

The positional fluctuations of Ca atoms (CA) ofEts-1 ETS domain evaluated from MD trajectoriesare shown in Figure 4 along with that from thecrystal structure. Although the magnitude of thefluctuations from X-ray and MD structures aredifferent, the fluctuation pattern is similar. Thehelix-3 region (residues 386–396) of the ETSdomain that recognizes the core DNA sequencehas smaller fluctuations, compared to the otherpart of the protein. On the other hand, the turnregion (379–384) between helix-2 and helix-3 andthe winged region (405–410) show larger fluctu-

ations. The steep peak observed in the C-terminalof helix-1 (348–353) of ETS–GGAA MD structureis likely due to the contact between the residues348–353 and helix-5 region (430–436). However, itdoes not seem to affect any other structural fea-tures of the ETS domain.

DNA structure of Ets-1 ETSdomain–DNA complex

DNA bending

The time-variation plots of DNA bending anglefor the ETS–GGAA and ETS–GGAG MD struc-tures are given in Figure 5(a). According to theliterature,14,28 the DNA bending angle is defined asshown in Figure 5(b). Large fluctuations arenoticed in the bending angle with average valueof 168 and 228 for ETS–GGAA (900–3480 ps) andETS–GGAG (900–3930 ps), respectively. In GGAAhelix, a relatively extended DNA structure isobtained compared to the ETS–DNA complexes.The average (900–3255 ps) bending angle ofGGAA is found to be 118. A stereo plot of averageDNA structures of ETS–GGAA (900–3480 ps),ETS–GGAG (900–3930 ps) and GGAA (900–3255 ps) are given in Figure 5(c)–(e), superimposedon the canonical B-DNA structure.29 The plots indi-cate that the presence of ETS domain influences theDNA bending. As can be seen in Figure 5(a), (c),and (d), no significant difference in DNA bending

Figure 2. Sequences of Ets-1 ETSdomain and 14 base-pair DNA: (a)amino acid sequence with the resi-due numbers and secondary struc-ture indicated above the sequence,(b) DNA base sequence with thenumbering provided above orbelow the sequence. Residues inthe core GGAA are shown in red.In the GGAG sequence, themutated GC base-pair is shown inblue.

MD Simulations of Ets-1 ETS Domain–DNA Complexes 347

is observed in the ETS–GGAA and ETS–GGAGcomplexes.

Major and minor groove widths

According to the literature,14 groove widths aredefined as distances between two appropriatephosphate atoms (see Method). The average majorand minor groove widths of 14 base-pair DNA inETS–GGAA, ETS–GGAG and GGAA are summar-ized in Table 1. The major groove width around thecore GGAA region is found to be comparativelylarge in the crystal structure (PDB code: 1K79),although the DNA bends into the major groove. Inthe MD averaged structure, an expansion of majorgroove width around the core region is alsoobserved. For example, the averaged major groovewidths at base-pair 1 in ETS–GGAA and GGAAare 20.43 A and 17.11 A, respectively. Thus, theMD simulation of the ETS domain–DNA complexreproduced this structural feature well. Note-worthy are the observations that fluctuations inthe major groove width around the core region(base-pairs 1–4) are quite small in ETS–GGAA,while large fluctuations are observed in the majorgroove width at the base-pairs 3 and 4 in ETS–GGAG complex. These small fluctuations in themajor groove width of ETS–GGAA may reflectthe stability of the interaction between the helix-3of ETS domain and the GGAA core sequence.

In comparison to the dynamics of the majorgroove, the minor groove widths obtained fromMD simulations of ETS–GGAA, ETS–GGAG andGGAA show little difference between each other.The values of minor groove width in the crystal

Figure 3. Time evolution of RMSD of the MD com-plexes of ETS–GGAA (black) and ETS–GGAG (red)with respect to the corresponding minimized structures:(a) the backbone heavy-atoms of the ETS domain and(b) all the heavy-atoms of DNA except terminal base-pairs. The RMSD value of the heavy-atoms of GGAAhelix (green) with respect to minimized structure isshown in (b).

Figure 4. Atomic fluctuations of Ca atoms of the Ets-1ETS domain in MD structures of ETS–GGAA (averagedfor 900–3480 ps) in black, and ETS–GGAG (averagedfor 900–3930 ps) in red. Fluctuations of Ca atom of thecrystal structure are shown in blue.

Table 1. Average major and minor groove widths (A)and their standard deviations (in parenthesis) of the 14base-pair DNA duplexes of ETS–GGAA, ETS–GGAGand GGAA MD structures. The values of X-ray structureof ETS–GGAA (PDB code: 1K79) are given

Base-pair

X-raystructure

(PDB:1K79)

ETS–GGAA(900–

3480 ps)

ETS–GGAG(900–

3930 ps)

GGAA(900–

3255 ps)

Major groove23 18.71 17.91(^1.50) 17.55(^1.64) 17.00(^1.49)22 17.90 17.91(^1.64) 18.29(^1.67) 17.54(^1.71)21 17.75 18.56(^1.17) 18.13(^1.15) 17.32(^1.55)1 20.47 20.43(^0.84) 19.33(^0.80) 17.11(^1.93)2 19.87 20.82(^0.43) 20.52(^0.40) 17.41(^1.78)3 18.45 20.40(^0.85) 19.10(^1.63) 16.80(^1.50)4 17.06 18.22(^0.75) 18.97(^1.79) 16.82(^1.24)5 18.82 17.94(^1.00) 19.96(^1.90) 15.90(^1.46)6 18.90 18.61(^1.54) 19.92(^1.85) 16.80(^1.40)

Minor groove23 13.88 14.20(^0.88) 13.86(^1.09) 13.70(^1.16)22 13.52 13.77(^1.27) 14.74(^1.21) 14.11(^1.39)21 14.90 15.48(^1.13) 15.81(^1.57) 14.57(^1.17)1 13.46 14.96(^1.05) 15.60(^1.28) 14.52(^1.14)2 14.14 14.60(^1.23) 14.68(^1.70) 13.84(^1.12)3 13.21 13.80(^1.24) 14.83(^1.33) 13.22(^0.88)4 9.98 12.67(^1.34) 13.54(^1.05) 13.11(^1.02)5 9.06 12.02(^1.21) 11.11(^1.17) 13.40(^1.02)


structure are consistent with those in the MD struc-ture of ETS–GGAA, except for the values at base-pairs 4 and 5.

Sugar puckering

The time evolution of pseudorotational phaseangles ðPÞ of selected nucleosides in both ETSdomain–DNA complexes are represented inFigure 6. The switching of sugar puckering fromC20-endo (S-form) to C30-endo (N-form) or from N-form to S-form is observed in some nucleoside resi-dues (Figure 6(b), (c), and (e)). On the other hand,the flexibility in sugar puckering is restricted insome DNA regions where the phosphates ornucleobases have contacts with the ETS domain.

Especially, the sugar conformations of G23 and A60

are found to be almost locked in S-form puckering(Figure 6(a) and (f)) due to their 30-phosphategroup being rigidly held in an interaction withspecific amino acid residues in the ETS domain.The 30-phosphate oxygen atoms, O1P and O2P ofG23 (by convention listed in Table 2 as the 50-phos-phate oxygen atoms of the neighboring C22,O1P(C22) and O2P(C22), respectively) contact withthe hydroxyl oxygen OH of Tyr386, side-chainamino nitrogen NZ of Lys404 and main-chainamino nitrogen N of Tyr410, while those of A60

(listed as O1P(T50) and O2P(T50) form a salt bridgewith N(Leu337) and OH(Tyr396). A considerabledifference in the puckering between the ETS–GGAA and ETS–GGAG structures is observed at

Figure 5. (a) Time evolution ofthe DNA bending angle u (deg) inthe MD structures of ETS–GGAA(black) and ETS–GGAG (red). Thebending angle of the crystal struc-ture is shown in blue. (b) Schematicrepresentation of definition of DNAbending angle u. Stereo view of theaveraged DNA structures obtainedfrom the MD simulations of (c)ETS–GGAA (900–3480 ps) inblack, (d) ETS–GGAG (900–3930 ps) in red and (e) GGAA(900–3255 ps) in green. The canoni-cal B-form DNA structure is alsoshown (blue) in (c)–(e) for compari-son. Superimpositioning is per-formed according to theorientations of base-pairs 24, 25and 26. All hydrogen atoms areomitted for clarification.


C21 (Figure 6(c)). This nucleoside residue in ETS–GGAA complex likely prefers the S-type sugar con-formation, while the sugar puckering of C21 inETS–GGAG remains N-type after 2500 ps.

Interaction between Ets-1 ETS domain and DNA

Contacts of Arg391 and Arg394 with nucleobasesin the core sequence

The distances between non-bonded entities inthe contact region of the ETS domain–DNA com-plexes determined by MD simulation and crystalstructures are summarized in Table 3. The hydro-gen bonding structures involving Arg391 and

Arg394 with nucleobases are shown in Figure 7.The time variations of selected heavy-atom non-bonded distances are shown in Figure 8. In theETS–GGAA MD structure, Arg391 of ETS is inbidentate contacts with G2 nucleobase by hydrogenbonds between secondary amino nitrogen NE ofArg391 and O6 oxygen of G2, and the otherbetween guanido nitrogen NH2 of Arg391 and N7nitrogen of G2 (Figure 7(a)). These two hydrogenbonds are maintained during the entire simulation(black line of Figure 8(a) and (c)). Such hydrogenbond interactions are observed between Arg394and G1 nucleobase (Figure 7(b)) for the durationafter 1700 ps in ETS–GGAA (black line of Figure8(d) and (f)). These separations are large for the

Figure 6. Time evolution of the pseudorotation phase angle P of sugar ring of nucleosides (a) G23, (b) C230, (c) C21, (d)G2, (e) C20 and (f) A60 in the MD structures of ETS–GGAA (black) and ETS–GGAG (red).

Table 2. Average non-bonded distances (A) and their standard deviations (in parenthesis) of the contact sites of Ets-1–DNA phosphate backbone of ETS–GGAA and ETS–GGAG MD structures. The values of X-ray structure of ETS–GGAA (PDB code: 1K79) are given

ETS–GGAA ETS–GGAG

Atoms X-ray structure (PDB: 1K79) (900–1650 ps) (1800–3480 ps) (900–2100 ps) (2700–3930 ps)

N(Leu337)· · ·O1P(T50) 2.80 2.94(^0.20)* 2.96(^0.27)* 3.00(^0.29)* 2.96(^0.18)*

NE1(Trp375)· · ·O2P(T40) 2.86 3.53(^0.50) 3.22(^0.51)** 2.96(^0.31)* 3.02(^0.40)*

NZ(Lys379)· · ·O1P(T40) 2.85 4.87(^1.18) 5.35(^0.67) 2.77(^0.18)* 2.73(^0.14)*

OH(Tyr386)· · ·O2P(C22) 2.52 2.65(^0.10)* 2.65(^0.10)* 2.65(^0.10)* 2.65(^0.10)*

NZ(Lys388)· · ·O2P(T30) 2.95 4.71(^0.32) 2.79(^0.33)* 2.75(^0.13)* 2.70(^0.10)*

OH(Tyr396)· · ·O2P(T50) 2.46 2.64(^0.10)* 2.81(^0.37)** 2.94(^0.49)** 2.64(^0.10)*

OH(Tyr397)· · ·O2P(C21) 2.66 2.74(^0.14)* 2.72(^0.13)* 2.71(^0.12)* 2.69(^0.12)*

NZ(Lys399)· · ·O2P(A60) 2.64 3.79(^1.21) 3.72(^0.93) 3.98(^0.94) 3.38(^0.83)NZ(Lys404)· · ·O1P(C22) 2.42 2.72(^0.11)** 2.74(^0.12)** 2.87(^0.33)** 2.73(^0.11)**

N(Tyr410)· · ·O1P(C22) 2.83 2.83(^0.13)* 2.87(^0.15)* 3.01(^0.28)* 3.00(^0.21)*

OH(Tyr410)· · ·O1P(G23) 2.58 2.86(^0.48)** 3.03(^0.69)** 2.66(^0.15)* 2.64(^0.10)*

Values within *0–0.25 A and **0.25–0.50 A different from the X-ray structure.


time-period 250–1700 ps, during which a hydro-gen bond interaction is observed between the gua-nido nitrogen NH2 of Arg394 and N7 nitrogen ofG1 nucleobase (black line of Figure 8(e)).

In the case of ETS–GGAG, the contact betweenNE of Arg391 and O6 of G2 (red line of Figure8(a)) give way to a hydrogen bond between NH2of Arg391 and O6 of G2 during 2850–2900 ps(Figure 7(c) and red line of Figure 8(b)). AlthoughNH2 of Arg391 is within 3.0 A distance from N7nitrogen of G2 even after 2850 ps, the non-bondedangle for NH2(Arg391)–HH21(Arg391)· · ·N7(G2)

is 120.7(^27.5)8. This does not allow hydrogenbond formation (Figure 7(c)).30 Arg394 has a con-tact with G1 forming two hydrogen bonds onebetween NE of Arg394 and N7 nitrogen of G1 andthe other between NH2 of Arg394 and O6 oxygenof G1. These interactions exist until 2100 ps ofdynamics (red line of Figure 8(d) and (f)). The con-tact between NH1 of Arg394 and OG of Ser390,which would stabilize the helix-3 structure, is alsobroken during the same time (data not shown). Inplace, NH2 of Arg394 forms a hydrogen bondwith N7 nitrogen of G1 (Figures 7(d) and 8(e)). So

Table 3. Average non-bonded distances (A) and their standard deviations (in parenthesis) of the contact sites of Ets-1–DNA nucleobase of ETS–GGAA and ETS–GGAG MD structures. The values of X-ray structure of ETS–GGAA (PDBcode: 1K79) are given

ETS–GGAA ETS–GGAG

Atoms X-ray structure (PDB: 1K79) (900–1650 ps) (1800–3480 ps) (900–2100 ps) (2700–3930 ps)

NE(Arg391)· · ·O6(G2) 2.65 2.90(^0.15)* 2.96(^0.19)** 2.97(^0.17)** 3.91(^0.46)NH2(Arg391)· · ·O6(G2) 3.67 3.94(^0.29)** 4.20(^0.35) 4.19(^0.31) 2.97(^0.33)NH2(Arg391)· · ·N7(G2) 2.77 2.90(^0.10)* 2.90(^0.13)* 2.89(^0.10)* 3.00(^0.15)*

NE(Arg394)· · ·N7(G1) 2.84 3.96(^0.53) 2.97(^0.12)* 2.94(^0.13)* 5.16(^0.17)NH2(Arg394)· · ·N7(G1) 3.67 3.04(^0.21) 3.58(^0.28)* 3.67(^0.30)* 3.00(^0.13)NH2(Arg394)· · ·O6(G1) 2.85 3.97(^0.62) 2.91(^0.17)* 2.91(^0.17)* 5.13(^0.20)OH(Tyr395)· · ·N6(A3) 3.10 3.42(^0.38)** 3.09(^0.26)* 3.29(^0.27)* 4.27(^0.63)OH(Tyr395)· · ·N6/O6(A4/G4) 3.64 3.98(^0.60)** 3.63(^0.40)* 4.36(^0.38) 4.64(^0.49)OH(Tyr395)· · ·O4/N4(T40/C40) 3.77 3.52(^0.55)* 3.58(^0.51)* 3.08(^0.25) 3.00(^0.18)CD2(Tyr395)· · ·C5M(T50) 3.41 3.96(^0.28) 3.92(^0.38) 3.97(^0.37) 3.93(^0.34)CE2(Tyr395)· · ·C5M(T50) 3.42 3.82(^0.30)** 4.05(^0.43) 3.87(^0.31)** 3.75(^0.27)**

Values within *0–0.25 A and **0.25–0.50 A different from the X-ray structure.

Figure 7. Molecular plot showing the contact interactions of MD structures. (a) Arg391 with G2C20 DNA base-pairand (b) Arg394 with G1C10 base-pair in ETS–GGAA, averaged for 1800–3480 ps; (c) Arg391 with G2C20 base-pair and(d) Arg394 with G1C10 base-pair in ETS–GGAG, averaged for 2700–3930 ps. See the stable bidentate hydrogen bondsin (a) and (b), while they are absent in (c) and (d).


during the course of dynamics certain structuralalignments prevail favoring some interactions atthe cost of others.

Contact of Tyr395 with the DNA

In the crystal structure of Ets-1 ETS domain–DNA complex, Tyr395 is proximal to A4 and T40

nucleobases in the major groove of the core region(Figure 1).5 As shown in Table 3, the hydroxylgroup of Tyr395 is at 3.09 and 3.63 A from the exo-cyclic amino nitrogen atoms N6 of A3 and A4,respectively in the MD structure of ETS–GGAA.This indicates that the hydroxyl group forms ahydrogen bond with N6 of A3, while it makes aweak contact at A4. The interaction between thehydroxyl group of Tyr395 and the carbonyl oxygenO4 of T40 is also observed. However, the close con-tact, in which the distance between the hydroxylgroup of Tyr395 and O4 of T40 is less than 3.2 A, isidentified only for short time-periods (1000–1200 ps and 2250–2600 ps). The steric hindrancecaused by the 5-methyl group of T40 likely preventshydrogen bonding between the O4 carbonyl oxy-gen of T40 and the hydroxyl group of Tyr395.During dynamics the delta carbon CD2 of Tyr395is at 3.92 A from the 5-methyl carbon C5M of T50,indicating hydrophobic interaction between thephenyl ring of Tyr395 and the 5-methyl group ofT50 (Table 3).

In the MD structure of ETS–GGAG, a contactbetween the hydroxyl group of Tyr395 and N6atom of A3 is observed until about 2800 ps(Table 3). The Tyr395 hydroxyl group forms ahydrogen bond with the 4-amino nitrogen N4 ofC40 nucleobase. Unlike the ETS–GGAA complexwhere the 5-methyl group of T4 prevents hydrogenbonding, the C40 in ETS–GGAG is in contact withTyr395. The hydrophobic interaction between C5M

of T50 and the phenyl ring of Tyr395 is also seen inthe ETS–GGAG complex.

Motion of the helix-3 on the interface betweenETS domain and DNA

In order to investigate the motion of the helix-3in the major groove of the DNA, the structureswere averaged at the intervals of 300 ps and ana-lyzed. Some structures are superimposed accord-ing to the orientations of G2, A3 and A4/G4 (Figure9). The stereo pictures indicate that the positionand motion of the helix-3 in each complex arequite different. In the ETS–GGAA complex, thehelix-3 is settled in the major groove of the consen-sus DNA sequence without significant positionalfluctuations. The main-chain of the helix-3 (CA, Cand N atoms) is almost at the same position duringthe entire duration of MD simulation (Figure 9(a)),and the contact residues, Arg391, Arg394 andTyr395 show no change in conformation (Figure9(b)). In addition, overall the helix-3 region is simi-lar in both X-ray and MD averaged structures (datanot shown).

On the contrary, a distinct motion of the helix-3region can be seen in the ETS–GGAG complex(Figure 9(c) and (d)). The main-chain of the helix-3gradually moves as the simulation proceeds. Forexample, the Ca atom of Tyr395 in the MD struc-ture averaged for 1200–1500 ps (yellow structurein Figure 9(d)) shows a movement of 4.11 A withrespect to the MD structure averaged for 3000–3300 ps (blue structure in Figure 9(d)).

Interaction between the ETS domain andphosphate backbone in the DNA

Eleven direct contacts between hydrogen donorsin the ETS domain and phosphate backbone in the

Figure 8. Time-dependent vari-ation of separations of (a)NE(Arg391)· · ·O6(G2),(b) NH2(Arg391)· · ·O6(G2),(c) NH2(Arg391)· · ·N7(G2),(d) NE(Arg394)· · ·N7(G1),(e) NH2(Arg394)· · ·N7(G1) and(f) NH2(Arg394)· · ·O6(G1) of ETS–GGAA (black) and ETS–GGAG(red) MD structures.


Figure 9. Stereo diagrams of the helix-3 in the major groove of core DNA sequence in (a),(b) ETS–GGAA and (c),(d)ETS–GGAG MD structure. The MD structures averaged for the period 1200–1500 ps (yellow), 1800–2100 ps (green),2400–2700 ps (gray) and 3000–3300 ps (blue) are superimposed according to the orientations of G2, A3 and A4/G4. In(a) and (c), the backbone atoms (CA, C and N) of binding site (residues 386–396) corresponds to the major grooveview of DNA (base-pairs 1–5). In (b) and (d), the close-view of (a) and (c) perpendicular to the helix axis are given,with only few important protein residues (Arg391, Arg394 and Tyr395) and nucleobases (G1, G2, A3, A4/G4, T40 andT50) shown.


DNA are observed in the crystal structure of theETS–GGAA complex (PDB code: 1K79).5 Theseare listed in Table 2, along with the correspondingdistances of ETS–DNA MD complexes. Only slightdifferences (,0.5 A) in the contact distances of saltbridge formations between the ETS domain andphosphate backbone of DNA are observedbetween the crystal and the averaged MD struc-tures. In ETS–GGAA, the contacts between the pri-mary amino nitrogen NZ of Lys379 and thephosphate oxygen O1P of T40 and between NZ ofLys399 and O2P of A6 are not seen.

As seen in Figure 8, the drastic changes areobserved in the hydrogen bonding patternbetween DNA base and the helix-3 region of theETS–GGAG MD structure. However, no consider-able change in the direct contact of the turn andwinged region of the ETS domain (two ends of thehelix-3 region) with the DNA phosphate backboneis noticed in the MD averaged structures (900–2100 and 2700–3930 ps) of ETS–GGAG. This resultsuggests the possibility that the helix-3 works inde-pendently of the flanked regions in the ETSdomain–DNA interaction.

Discussion

Comparison of the MD structures to theexperimental data

MD simulations on ETS–GGAA and ETS–GGAG clearly show the presence of meta-stablestates of hydrogen bonding in which the conservedresidues, Arg391 and Arg394, are participating(Figure 7). In the low affinity ETS–GGAG complex,both arginine residues change hydrogen bondingpartners after 2 ns MD simulation. In contrast,Arg394 in ETS–GGAA changes only its side-chainconformation preference at around 1.7 ns to formmore stable bidentate hydrogen bonds with thesame partner (Figure 8(d) and (f)). Thus, the argi-nine residues show a certain degree of confor-mational flexibility in the ETS domain–DNAcomplexes. These observations agree well with theresults from NMR experiments of Ets-112,13 andFli-1,11 in which the conserved arginine residues inthe ETS domain–DNA complexes were notassigned, and it was concluded that the Arg391and Arg394 did not have a single defined confor-mation in the complexes. Furthermore, the hydro-gen bonding mode of these arginine residues inthe crystal structures of ETS domain–DNA com-plexes depends on the complex studied,5 – 10 whichalso supports the conformational flexibility of thearginine residues.

In the crystal structures of ETS domain–DNAcomplexes, the DNA bending angle varies fromstructure to structure.5 – 10 The X-ray structure ofPU.1–DNA complex shows a DNA bending angleof 88,9,10 while the value in the Ets-1–DNA complexwas reported5 to be 278. MD studies on ETS–GGAA and ETS–GGAG show major fluctuations

in the bending angle in the range 5–398 and 5–428, respectively (Figure 5(a)). These results indi-cate that the differences in the DNA bendingangle are likely to arise due to flexibility of DNAhelix in the complexes.

Analysis of Ets-1 ETS domain–DNA complex inX-ray structures has led to the proposal that thereduction in binding affinity of ETS–GGAG com-plex is due to the absence of van der Waals con-tacts with C40, and the reduction in the van derWaals overlap between the 5-methyl group of T50

and the phenyl ring of Tyr395.5 The possibility ofhydrogen bond formation between the hydroxylgroup in Tyr395 and N4 of C40 was also reportedin the literature.5 However, it was concluded thatthe contact would not significantly affect the bind-ing energy. These explanations from X-ray analysisare not consistent with the MD simulations. Thedistance between the 5-methyl group of T50 andthe delta carbon CD2 of Tyr395 observed in theMD structure of ETS–GGAA are comparable tothat of ETS–GGAG (Table 3), indicating that thevan der Waals interaction between the 5-methylgroup of T50 and the phenyl ring of Tyr395 doesnot play an important role in the recognition ofthe GGAA core sequence. In addition, the hydroxylgroup of Tyr395 is found to be hydrogen bonded toN4 of C40 in the MD simulation of ETS–GGAG andis maintained during the entire simulation. How-ever, in ETS–GGAA the contact of the hydroxylgroup with O4 of T40 is only observed intermit-tently. These results indicate the significance of thecontact between the hydroxyl group of Tyr395 andN4 of C40. It is noteworthy that the specific recog-nition of the nucleobase 40-position by ETS domainis observed in ETS–GGAG, but not in ETS–GGAA.

Direct and indirect readout mechanism

X-ray crystallographic analysis of protein–DNAcomplexes often reveals a distorted structure ofDNA helix comprising a bending and a kinkingstructure.31,32 In some of these cases, it seems to bedifficult to explain all the sequence-specificitybased only by the direct and water-mediated inter-actions between the protein and such a distortedDNA. An indirect readout mechanism has beenproposed to explain the sequence-specific DNAbinding of protein.33 – 37 In the indirect readoutmechanism, a protein recognizes a sequence-dependent DNA conformation that already existsbefore binding or, alternatively, is induced afterbinding. A comparative analysis of the DNA bind-ing specificity of other ETS family proteins (Fli-1,SAP-1, PU.1 and TEL) using a multiplex and otherexperimental techniques was reported, wherein apossibility of the indirect readout mechanism inrecognition of GGA core flanking regions (23,22, 21, 4, 5 and 6 base-pairs) was mentioned.38

However, no significant difference in either DNAbending angle (Figure 5(a), (d), and (e)) and ETSdomain–DNA phosphates interactions (Table 3) areobserved between ETS–GGAA and ETS–GGAG


during MD simulations. This is sufficient toexplain an advanced stability of the ETS–GGAAcomplex. These results suggest that the AT base-pair at position þ4 is recognized by the directreadout mechanism, but not by the indirect read-out mechanism in the Ets-1 ETS domain–DNAcomplex.

Role of Arg391, Arg394 and Tyr395 inrecognition of the core DNA sequence

The MD studies on the ETS–GGAA and ETS–GGAG complexes clearly show that the highly con-served Arg391 and Arg394 play an important rolein the interaction between the helix-3 and themajor groove of GGAA core sequence (Figures 7and 8) consistent to X-ray studies of ETS domain–DNA complexes.5 – 10 The two bidentate hydrogenbonds formed between Arg391 and G2, andbetween Arg391 and G1 are stable in the MD simu-lation of ETS–GGAA, while the correspondinginteractions are not maintained in the simulationof ETS–GGAG. The decrease in the stability of theETS–GGAG complex should be due to the collapseof these bidentate interactions. It is of great interestthat the two arginine residues, Arg391 and Arg394,show such a different behavior in the high and lowaffinity complexes, though they contact the con-served GG sequence (þ1 and þ2 positions) ineach complex. These arginine residues are highlyconserved among the ETS protein family and arewell known to take part in DNA binding. Thus,the results obtained here are likely to be meaning-ful for a clear understanding of the sequence-specific DNA binding of the ETS domain.

The Tyr395 residue of the ETS domain plays acritical role in the recognition of the GG sequenceby the conserved arginine residues, Arg391 andArg394. Tyr395 is a neighboring residue ofArg394. Therefore, the motion of Tyr395 shouldhave a direct influence on the location or move-ment of Arg394. In the MD structure of ETS–GGAG, the hydroxyl group of Tyr395 is hydrogenbonded with N4 of C40 at the beginning of simu-lation. This hydrogen bond formation resulted, tosome extent, in the motion of Tyr395 in the majorgroove of the GGAG sequence. The Arg394 isdirectly affected by the motion of Tyr395 and thebidentate hydrogen bonds between Arg394 and G1

are lost at 2100 ps (Figure 8(a) and (c)). Thereafter,a large amount of motion of the helix-3 in themajor groove of GGAG sequence is observed(gray structure of Figure 9(c) and (d)). As a resultof these motions, the bidentate interactionsbetween the Arg391 and G2 are absent after2850 ps. These conformational changes are relatedto each other and arise from hydrogen bond for-mation between the hydroxyl group of Tyr395 andN4 of C40.

On the contrary, the helix-3 region is localized inthe major groove of the core DNA sequence ofETS–GGAA without considerable fluctuations,and consequently the conserved arginine residues

form stable hydrogen bonds in the MD structure(Figure 9). Unlike the ETS–GGAG complex, thehydroxyl group of Tyr395 does not have a stableinteraction with O4 of T40 due to a steric hindranceby the 5-methyl group of T40. Instead, the hydroxylgroup makes a contact with N6 of A3. Conse-quently, the position of Tyr395 and the helix-3does not change during the MD simulation, allow-ing a stable interaction between the helix-3 andthe GGAA core sequence. Thus, the Tyr395 mightwork as a tactile sensor to distinguish a targetedAT base-pair at the þ4 position.

Conclusions

We conducted 3.5–3.9 ns MD simulations of Ets-1 ETS domain–14 base-pair DNA complexes withPME treatment of electrostatic interactions. TheseMD simulations have provided us a good deal ofinformation on the sequence specific interactionbetween ETS domain and the consensus DNA, asschematically shown in Figure 10. Two conservedarginine residues, Arg391 and Arg394, play animportant role in binding with the GGAA coresequence. Although these arginine residues showcertain flexibility in side-chain conformation, theymake bidentate contacts with G1 and G2 to stabilizethe complex structure. The contacts between thesearginine residues and GG dinucleotides in thecore sequence are regulated by the motion ofTyr395. In the high-affinity complex ETS–GGAA,the hydrogen bonding between the hydroxylgroup of Tyr395 and the 4-carbonyl oxygen of T40

is prevented by the bulky 5-methyl group of T40.Rather, Tyr395 makes a contact with A3, allowinghelix-3 to be in the appropriate location in themajor groove of DNA. On the contrary, thehydroxyl group of Tyr395 is hydrogen bondedwith the 4-nitrogen of C40 in the low affinity com-plex ETS–GGAG. This hydrogen bonding causesmotion in the helix-3 region and results in the dis-ruption of bidentate contacts between the con-served arginine residues and the G1G2

dinucleotides. Thus Arg391, Arg394 and Tyr395 inhelix-3 of Ets-1 work cooperatively to recognizethe GGAA core sequence.

Methods

Modeling of initial structures

The crystal structure (PDB code: 1K79) of Ets-1 ETSdomain–15 base-pair DNA complex5 was used for prep-aration of the starting structure of ETS–GGAA.Although the crystal structure involved two complexesin the asymmetric unit, these two structures are essen-tially identical. So, the first complex was considered inour study. In the crystal structure, the 15 base-pair DNAhas a 50-overhang structure, so the 50-terminal nucleotidein each strand was deleted. An unusual hydrogen bondpattern was observed at base-pair 14 in the crystal struc-ture. The glycosidic bond angle (x) at A14 was adjusted to


allow the base-pair 14 to form Watson–Crick type hydro-gen bonds using the program INSIGHT II (version 97).The initial structure of ETS–GGAG was built by repla-cing the AT base-pair at position þ4 in ETS–GGAAwith GC base-pair (Figure 2) using the INSIGHT II pack-age so that all the conformations of other residues are thesame. The initial structure of GGAA (the 14 base-pairDNA alone) was prepared by removing the ETS domainfrom the initial structure of the ETS–GGAA complex.

The hydrogen atoms were added using HBUILD ofthe CHARMM program.39 On the basis of protonationsites, the imidazoles of His403 and His430 of Ets-1 areprotonated at both ND1 and NE2 positions in the ETSdomain–DNA complexes. The ETS domain has a chargeof þ6, while the 14 base-pair DNA includes a charge of226. To neutralize the net charge of each ETS–DNAcomplex, appropriate ions were placed near the phos-phate oxygen atoms of the DNA and also near the sol-vent-exposed charged residues of Ets-1. For the

simulation on GGAA, 26 Naþ were added 3.5 A awayfrom the phosphorous atom in each strand of the duplex.Then, each system was minimized for 50 steps with stee-pest descent (SD) method.

Molecular simulations

MD Simulations were performed using the programCHARMM (version c27b4)40 with all-atom force fieldparameters.41 Periodic boundary conditions were definedusing an orthorhombic box of dimensions67.4 A £ 62.7 A £ 54.3 A for ETS–GGAA and ETS–GGAG structures, and 45.8 A £ 63.8 A £ 44.8 A forGGAA filled with TIP3P42 model water molecules. Thewater molecules in the box were minimized for 100steps of SD method and equilibrated for a period of30 ps constant pressure–temperature (NPT) dynamics.Then the water box was overlaid onto the Ets-1 ETS

Figure 10. Schematic diagrams of the hydrogen bonds between helix-3 and core DNA sequence in (a) ETS–GGAAand (b) ETS–GGAG complexes.


domain–DNA complex with ions and crystal waters.Solvent molecules with oxygen atoms within 1.6 A ofnon-hydrogen atoms in the DNA helices and thosewithin 2.5 A of any other non-hydrogen atoms weredeleted. The total number of atoms was 24,959 for ETS–GGAA, 24,961 for ETS–GGAG and 14,479 for GGAA.Positions of water molecules were minimized for 100steps of SD followed by 400 steps of adopted basis New-ton–Raphson (ABNR) methods in each structure, keep-ing all solute molecules fixed. After that, the constraintson ions were released, and the minimization for 100steps of SD followed by 400 steps of ABNR methodswere performed. Then, the entire system was minimizedfor 100 steps with SD and 2000 steps with ABNRmethods before starting simulations.

Leapfrog Verlet integration scheme43 was used with anintegration time-step of 1.5 fs. SHAKE44 was applied toall covalent bonds involving hydrogen atoms. Imageswere generated using the CRYSTAL module ofCHARMM. A constant dielectric of unity was used. Elec-trostatic interactions were treated with PMEformalism45,46 as implemented47 in the CHARMM pro-gram. PME calculations were performed using realspace cutoff of 10 A with Lennard–Jones interactionstruncated at the same distance. A convergence parameter(k) of 0.36 A21 and a sixth degree B-spline interpolationwere used with the PME method.

During the equilibration, the structure was relaxed instages, so that the most strained parts of the systemcould adjust without artifacts. Initially, harmonic con-straint of 100 kcal mol2 A22 was applied to atoms otherthan waters and 21 ps simulation was performed at298 K. Then, the constraints on ions were released andthe system was heated gradually from 0 K to 298 K, atincrements of 100 K, each for 21 ps. Next, all the con-straints on solute molecules were removed and NOE(nuclear overhauser effect)-like distance constraintswere applied on the Watson–Crick hydrogen bonds atthe 30 and 50 end base-pairs of the DNA to reduce theend effects of DNA. Then, the system was re-equilibratedby heating the entire model at increments of 50 K for21 ps each from 0 K to 298 K. These stages were carriedout in NPT ensemble (with temperature and pressure of298 K and 1 atm, respectively), so that the water boxcould equilibrate in accord with the number of watermolecules. The dimensions of water box were allowedto vary in all directions. For subsequent simulations, theconstant volume–temperature ensemble was used, as itprovides more stable trajectories.48 Then, an additional30 ps of simulation was run at 298 K, to equilibrate theentire system at this temperature. The heating and equili-bration phases of dynamics lasted a total period of 240 psfor each system. The production simulation was thencontinued, at an average temperature of 298 K. Thesimulations for ETS–GGAA and ETS–GGAG were per-formed for total durations of 3.5 ns and 3.9 ns, respect-ively, while the simulation for GGAA helix was carriedout for 3.3 ns.

Structural analysis

The RMSD values of the Ets-1 ETS domain were eval-uated by least square fitting the backbone heavy-atomsto the minimized structure, while for DNA helices allthe heavy-atoms excluding the end base-pairs were con-sidered. The positional fluctuations of Ca in the ETSdomain backbone were calculated from MD trajectory,averaged over the period 900–3480 ps in ETS–GGAA

complex, and 900–3930 ps in ETS–GGAG. The exper-imental positional fluctuations were obtained using theequation ðDr2Þ1=2 ¼ ð3B=8p2Þ1=2 from Debye–WallerB-factors of Ets-1 ETS domain–DNA complex solved at2.4 A resolution (PDB code: 1K79).5 The averaged struc-ture was obtained by least squares fitting of all theatoms of the complex saved at 0.75 ps interval from thetrajectories to the minimized structure. Such averagedstructures were minimized for 500 steps of SD for mol-ecular plots, drawn using the programs MOLSCRIPT,49

Raster3D50 and MIDAS PLUS.51,52

DNA bending angle u, defined as the angle betweennormal vector of base-pair 22 and that of base-pair614,28 (Figure 5(a)) was evaluated using the programFreehelix98.31,32 According to the literature,14 the majorand minor groove widths were defined as the distancesbetween phosphorous atoms in different strands separ-ated by 3–4 base-pairs, i.e. P0ði 2 2Þ· · ·Pði þ 2Þ across theminor groove with three intervening base-pairs andPði 2 2Þ· · ·P0ði þ 2Þ across the major groove with fourintervening base-pairs. Here, PðiÞ and P0ðiÞ are the 50-phosphates of the complementary nucleotides that com-prise base-pair i; with the prime used to denote thecomplementary strand.

Acknowledgements

We thank Sun Hur (UCSB) for helpful discus-sions. This work was supported by NIH grant5R37DK0917136. We acknowledge computer timeon UCSB’s SGI Origin 2000.

References

1. Lelievre, E., Lionneton, F., Soncin, F. & Vandenbunder,B. (2001). The Ets family contains transcriptional acti-vators and repressors involved in angiogenesis. Int.J. Biochem. Cell. Biol. 33, 391–407.

2. Sharrocks, A. D., Brown, A. L., Ling, Y. & Yates, P. R.(1997). The ETS-domain transcription factor family.Int. J. Biochem. Cell. Biol. 29, 1371–1387.

3. Wasylyk, B., Hahn, S. J. L. & Giovane, A. (1993). TheEts family of transcription factors. Eur. J. Biochem.211, 7–18.

4. Dittmer, J. & Nordheim, A. (1998). Ets transcriptionfactors and human disease. Biochim. Biophys. ActaRev. Cancer, 1377, F1–F11.

5. Garvie, C. W., Hagman, J. & Wolberger, C. (2001).Structural studies of Ets-1/Pax5 complex formationon DNA. Mol. Cell, 8, 1267–1276.

6. Mo, Y., Vaessen, B., Johnston, K. & Marmorstein, R.(2000). Structure of the Elk-1–DNA complex revealshow DNA-distal residues affect ETS domain recog-nition of DNA. Nature Struct. Biol. 7, 292–297.

7. Mo, Y., Vaessen, B., Johnston, K. & Marmorstein, R.(1998). Structures of SAP-1 bound to DNA targetsfrom the E74 and c-fos promoters: Insights intoDNA sequence discrimination by ETS proteins. Mol.Cell, 2, 201–212.

8. Batchelor, A. H., Piper, D. E., de la Brousse, F. C.,McKnight, S. L. & Wolberger, C. (1998). The structureof GABP alpha/beta: an ETS domain ankyrin repeatheterodimer bound to DNA. Science, 279, 1037–1041.

9. Kodandapani, R., Pio, F., Ni, C. Z., Piccialli, G.,


Klemsz, M., McKercher, S. et al. (1996). A new pat-tern for helix-turn-helix recognition revealed by thePU.1 ETS-domain–DNA complex. Nature, 380,456–460.

10. Pio, F., Kodandapani, R., Ni, C.-Z., Shepard, W.,Klemsz, M., McKercher, S. R., Maki, R. A. & Ely,K. R. (1996). New insights on DNA recognition byets proteins from the crystal structure of the PU.1ETS domain–DNA complex. J. Biol. Chem. 271,23329–23337.

11. Liang, H., Mao, X. H., Olejniczak, E. T., Nettesheim,D. G., Yu, L. P., Meadows, R. P. et al. (1994). Solutionstructure of the Ets domain of Fli-1 when bound toDNA. Nature Struct. Biol. 1, 871–876.

12. Werner, M. H., Clore, G. M., Fisher, C. L., Fisher, R. J.,Trinh, L., Shiloach, J. & Gronenborn, A. M. (1995).The solution structure of the human Ets1–DNAcomplex reveals a novel mode of binding and trueside-chain intercalation. Cell, 83, 761–771.

13. Werner, M. H., Clore, G. M., Fisher, C. L., Fisher, R. J.,Trinh, L., Shiloach, J. & Gronenborn, A. M. (1997).Correction of the NMR structure of the ETS1/DNAcomplex. J. Biomol. NMR, 10, 317–328.

14. Kosikov, K. M., Gorin, A. A., Lu, X.-J., Olson, W. K. &Manning, G. S. (2002). Bending of DNA by asym-metric charge neutralization: all-atom energy simu-lations. J. Am. Chem. Soc. 124, 4838–4847.

15. Strauss, J. K. & Maher, L. J. (1994). DNA bending byasymmetric phosphate neutralization. Science, 266,1829–1834.

16. StraussSoukup, J. K. & Maher, L. J. (1997). Role ofasymmetric phosphate neutralization in DNA bend-ing by PU.1. J. Biol. Chem. 272, 31570–31575.

17. Cheatham, T. E., III, Cieplak, P. & Kollman, P. A.(1999). A modified version of the Cornell et al. Forcefield with improved sugar pucker phases and helicalrepeat. J. Biomol. Struct. Dynam. 16, 845–862.

18. Foloppe, N. & MacKerell, A. D., Jr (2000). All-atomempirical force field for nucleic acids: I. Parameteroptimization based on small molecule and con-densed phase macromolecular target data. J. Comput.Chem. 21, 86–104.

19. MacKerell, A. D., Jr & Banavali, N. K. (2000). All-atom empirical force field for nucleic acids: II. Appli-cation to molecular simulations of DNA and RNA insolution. J. Comput. Chem. 21, 105–120.

20. Scott, W. R. P., Hunenberger, P. H., Tironi, I. G., Mark,A. E., Billeter, S. R., Fennen, J. et al. (1999). The GRO-MOS biomolecular simulation program package.J. Phys. Chem. A, 103, 3596–3607.

21. Sagui, C. & Darden, T. A. (1999). Molecular dynamicssimulations of biomolecules: long-range electrostaticeffects. Annu. Rev. Biophys. Biomol. Struct. 28,155–179.

22. Giudice, E. & Lavery, R. (2002). Simulations ofnucleic acids and their complexes. Accts Chem. Res.35, 350–357.

23. Hagman, J. & Grosschedl, R. (1992). An inhibitorycarboxyl-terminal domain in Ets-1 and Ets-2 med-iates different binding of ETS family factors to pro-moter sequences of the mb-1 gene. Proc. Natl Acad.Sci. USA, 89, 8889–8893.

24. Jonsen, M. D., Petersen, J. M., Xu, Q. P. & Graves, B. J.(1996). Characterization of the cooperative functionof inhibitory sequences in Ets-1. Mol. Cell. Biol. 16,2065–2073.

25. Lim, F., Kraut, N., Frampton, J. & Graf, T. (1992).DNA-binding by c-Ets-1, but not v-Ets, is repressed

by an intramolecular mechanism. EMBO J. 11,643–652.

26. Petersen, J. M., Skalicky, J. J., Donaldson, L. W.,McIntosh, L. P., Alber, T. & Graves, B. J. (1995).Modulation of transcription factor Ets-1 DNA bind-ing: DNA-induced unfolding of an alpha helix.Science, 269, 1866–1869.

27. Wasylyk, C., Kerckaert, J. P. & Wasylyk, B. (1992). Anovel modulator domain of Ets transcription factors.Genes Dev. 6, 965–974.

28. Zhurkin, V., Lysov, Y. & Ivanov, V. (1979). Anisotro-pic flexibility of DNA and the nucleosomal structure.Nucl. Acids Res. 6, 1081–1096.

29. Arnott, S. & Hukins, D. W. (1972). Optimised par-ameters for A-DNA and B-DNA. Biochem. Biophys.Res. Commun. 47, 1504–1509.

30. Berendsen, H. J., van Gunsteren, W. F., Zwinderman,H. R. & Geurtsen, R. G. (1986). Simulations of pro-teins in water. Annu. N.Y. Acad. Sci. 482, 269–286.

31. Dickerson, R. E. & Chiu, T. K. (1997). Helix bendingas a factor in protein/DNA recognition. Biopolymers,44, 361–403.

32. Dickerson, R. E. (1998). DNA bending: the preva-lence of kinkiness and the virtues of normality. Nucl.Acids Res. 26, 1906–1926.

33. Wellenzohn, B., Flader, W., Winger, R. H.,Hallbrucker, A., Mayer, E. & Liedl, K. R. (2002).Indirect readout of the trp-repressor-operator com-plex by B-DNA’s backbone conformation transitions.Biochemistry, 41, 4088–4095.

34. Horton, N. C., Dorner, L. F. & Perona, J. J. (2002).Sequence selectivity and degeneracy of a restrictionendonuclease mediated by DNA intercalation.Nature Struct. Biol. 9, 42–47.

35. Chen, S. F., Vojtechovsky, J., Parkinson, G. N.,Ebright, R. H. & Berman, H. M. (2001). Indirect read-out of DNA sequence at the primary-kink site in theCAP–DNA complex: DNA binding specificitybased on energetics of DNA kinking. J. Mol. Biol.314, 63–74.

36. Chen, S. F., Gunasekera, A., Zhang, X. P., Kunkel,T. A., Ebright, R. H. & Berman, H. M. (2001). Indirectreadout of DNA sequence at the primary-kink site inthe CAP–DNA complex: alteration of DNA bindingspecificity through alteration of DNA kinking. J. Mol.Biol. 314, 75–82.

37. Martin, A. M., Sam, M. D., Reich, N. O. & Perona, J. J.(1999). Structural and energetic origins of indirectreadout in site-specific DNA cleavage by a restrictionendonuclease. Nature Struct. Biol. 6, 269–277.

38. Szymczyna, B. R. & Arrowsmith, C. H. (2000). DNABinding specificity studies of four ETS proteins sup-port an indirect read-out mechanism of protein–DNA recognition. J. Biol. Chem. 275, 28363–28370.

39. Brunger, A. & Karplus, M. (1988). Polar hydrogenpositions in proteins: empirical energy placementand neutron diffraction comparison. Protein: Struct.Funct. Genet. 4, 148–156.

40. Brooks, B. R., Bruccoleri, R. E., Olafson, B. D., States,D. J., Swaminathan, S. & Karplus, M. (1983).CHARMM: a program for macromolecular energy,minimization, and dynamics calculations. J. Comput.Chem. 4, 187–217.

41. MacKerell, A. D., Jr, Bashford, D., Bellott, M., Dun-brack, R. L., Evanseck, J. D., Field, M. J. et al. (1998).All-atom empirical potential for molecular modelingand dynamics studies of proteins. J. Phys. Chem. B,102, 3586–3616.

42. Jorgensen, W. L., Chandrasekhar, J., Madura, J. D.,


Impey, R. W. & Klein, M. L. (1983). Comparison ofsimple potential functions for simulating liquidwater. J. Chem. Phys. 79, 926–935.

43. Verlet, L. (1967). Computer “experiments” on classi-cal fluids. I. Thermodynamical properties of Len-nard–Jones molecules. Phys. Rev. 159, 98–103.

44. Ryckaert, J. P., Ciccotti, G. & Berendsen, H. J. C.(1977). Numerical integration of the Cartesianequations of motion of a system with constraints:molecular dynamics of n-alkanes. J. Comput. Phys.23, 237–241.

45. Darden, T., York, D. & Pedersen, L. (1993). Particlemesh Ewald: an N* logðNÞ method for Ewald sumsin large systems. J. Chem. Phys. 98, 10089–10092.

46. Petersen, H. G. (1995). Accuracy and efficiency of theparticle mesh Ewald Method. J. Chem. Phys. 103,3668–3679.

47. Feller, S. E., Pastor, R. W., Rojnuckarin, A., Bogusz, S.& Brooks, B. R. (1996). Effect of electrostatic forcetruncation on interfacial and transport properties ofwater. J. Phys. Chem. 100, 17011–17020.

48. Brown, D. & Clarke, J. H. R. (1984). A Comparison ofconstant energy. Constant temperature and constantpressure ensembles in molecular-dynamics simu-lations of atomic liquids. Mol. Phys. 51, 1243–1252.

49. Kraulis, P. J. (1991). MOLSCRIPT: a program to pro-duce both detailed and schematic plots of proteinstructures. J. Appl. Crystallog. 24, 945–949.

50. Merritt, E. A. & Bacon, D. J. (1997). Raster 3D: photo-realistic molecular graphics. In Macromolecular

Crystallography, Part B, Methods in Enzymology,vol. 277, pp. 505–524, Academic Press, San Diego,CA.

51. Ferrin, T. E., Huang, C. C., Jarvis, L. E. & Langridge,R. (1988). The Midas display system. J. Mol. Graph.6, 13–27.

52. Huang, C. C., Pettersen, E. F., Klein, T. E., Ferrin, T. E.& Langridge, R. (1991). Conic—a fast renderer forspace-filling molecules with shadows. J. Mol. Graph.9, 230–236.

Edited by B. Honig

(Received 14 February 2003; received in revised form5 May 2003; accepted 3 June 2003)

Supplementary Material for this paper compris-ing one Table and one Figure is available onScience Direct


sequence specific dna binding of ets-1 transcription factor

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