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Page 1: Supporting Information - PNAS...May 13, 2010  · as described previously (15), and analyzed using TENSOR2 (16). NMR-monitored titrations were carried out by recording sen- sitivity

Supporting InformationNelson et al. 10.1073/pnas.0915137107SI TextSI Materials and Methods. In vivo transcription assays and immuno-precipitation. Transient expression assays were performed aspreviously described in NIH3T3 mouse fibroblasts (1). In brief,transfected plasmids included 2.5 μg of firefly luciferase reportervector driven by an artificial double ETS site, 0.1 μg of wild-typeor mutant FLAG-Ets1 expression vector (or empty vector), 0.1 μgconstitutively active MEK1 (2) or empty vector, and 1 μg Renillaluciferase expression vector pRL-null (Promega). After transfec-tion, cells were serum-starved to reduce constitutive signaling.Relative luciferase activity was determined as the ratio offirefly/Renilla activity. For expression controls and phosphoryla-tion status, ∼1.5 × 106 cells were transfected with 3 μg offull-length FLAG-Ets1 expression plasmid, 3 μg constitutivelyactive MEK1, and 50 μl lipofectamine (Invitrogen). As describedpreviously (1), antibodies against the FLAG tag were used forimmunoprecipitation, followed by immunoblot detection witheither anti-Ets1 antisera (UT2) (3) or phospho-specific antiseraagainst pThr38-Ets1 (Biosource).

Protein purification. Samples of unlabeled and uniformly 15N- or15N∕13C-labeled murine Ets1/2 constructs (Ets11-138;S26A,Ets129-138, Ets129-138;L49R, Ets129-138;D123R, Ets129-138;E127R,Ets142-138, Ets151-138, Ets11-52, Ets21-172, and Ets263-172) wereprepared as described previously (2, 4–6). Deletions were madeby PCR cloning and mutations introduced by the Quickchangemethod. The His6-tagged constructs were incubated with throm-bin (Roche) in 20 mM Tris pH 8.4, 150 mM NaCl, and 2.5 mMCaCl2 overnight. Thrombin and the cleaved His6-tag wereremoved using ρ-aminobenzamidine beads (Sigma) and TALONmetal affinity resin (BD Biosciences). Ets11-138 and Ets129-138were phosphorylated in vitro using 50 mM ATP and a 1∶20molarratio of ERK2∶Ets1 (1). After incubation at 30 °C for 1.5 hr, theactive bacterially expressed His6-tagged ERK2 was removed witha HisTrap FF column (GE Healthcare). Ets1 samples of differingphosphorylation states were separated on a Mono Q column (GEHealthcare) using a 0–400 mM KCl linear gradient in 20 mM TrispH 7.5, 10% glycerol, and 2 mMDTT buffer, followed by passagethrough a Superdex 75 column (GE Healthcare) equilibrated in50 mM Tris pH 7.5, 10% glycerol, 50 mM KCl, 0.1 mM EDTA,and 2 mM DTT. Phosphorylation states were verified by electro-spray ionization mass spectrometry (ESI-MS).

The gene encoding TAZ1 (CBP340-439) was cloned from themurine CBP gene (GenBank accession no. 70995311) into thepET28b (Novagen) vector for expression as a His-tagged proteinin Escherichia coli BL21ðλDE3Þ cells. The protein was preparedas described previously (7).

Sample concentrations were determined from predicted ε280absorbance values.

Isothermal titration calorimetry. Isothermal titration calorimetry(ITC) experiments were conducted on a VP-ITC Microcalori-meter , MicroCal LLC. To block reac t i ve cys te ines ,Ets11-138;S26A and 2P-Ets11-138;S26A were treated with 50 mMiodoacetamide at pH 8.0 and 25 °C for 1 hr, followed by quench-ing with 150 mM DTT at 25 °C for 30 min. Mass spectrometry(ESI-MS Quatro-II) revealed three acetamide modifications,on four possible cysteines, in both species. TAZ1 and themodified Ets11-138;S26A samples were equilibrated into 25 mMTris pH 7.9, 50 mM KCl by dialysis. After equilibration of theVP-ITC to 15.0 °C for each experiment, 7.5 μL aliquots of500 μMTAZ1 were titrated into 1.4 mL of 10 μM nonphosphory-

lated or dual phosphorylated Ets11-138;S26A. Data were analyzedwith Origin 7.0 MicroCal LLC software.

Although Ser26, which has been proposed as a potential Ets1phosphorylation site (8), was not found to be modified in thesestudies, the S26A mutation was introduced into an Ets11-138construct that was used to produce protein for ITC experimentsto avoid any potential complications.

Trypsin proteolysis. His6-FLAG-HMK tagged Ets11-138 was phos-phorylated as described above or mock-treated in the absence ofATP. Phosphorylated or mock-treated proteins (240 pmol) weretreated with 0–2 μg trypsin in 25 mM Tris pH 7.9, 1 mMDTT, and10 mM CaCl2. After incubation for 2 min at room temperature,the reactions were stopped by boiling in 3×SDS sample buffer for5 min. Samples were run on 15% SDS PAGE to resolve thedigested peptides. Trypsin digestions were arrested with 1%trifluoroacetic acid for analysis of peptides by mass spectrometry(ESI-MS, Quatro-II).

Circular dichroism spectroscopy. Protein samples were dialyzed into25 mM Tris pH 7.5, 50 mM KCl, and 2 mM DTT. Data werecollected on an AVIV Circular Dichroism SpectrophotometerModel 410 (Biomedical Inc). Mean residue molar ellipticitywas calculated as ½θ� ¼ 3;298ðAL − ARÞ∕ð½Ets1� · l · NAAÞ, whereAL and AR are the absorbance of left and right circularly polar-ized light, respectively, [Ets1] is the concentration of proteinin μM, l is the path length (1 cm) of the quartz cuvette, andNAA ¼ 158 is the number of amino acids. Thermal denaturationcurves were generated from measurements taken at 222 nm in2-°C increments between 40 °C and 94 °C with 2- to 3-min equili-bration time and 30-sec acquisition time. Midpoint Tm valueswere determined from fitting to a standard two-state proteinunfolding equation (9).

NMR spectroscopy.NMR spectra of the Ets1/2 and CBP fragmentswere recorded using Varian 500-MHz Unity and 600-MHz Inovaspectrometers and analyzed using NMRpipe (10) and Sparky(11). 1H, 13C, and 15N resonance assignments were obtainedvia standard heteronuclear correlation experiments (12), asdescribed previously (4, 5, 13), initially using Ets 1/2 proteinsin 20 mM sodium phosphate (pH 6.2 for Ets129-138 and pH 6.3for 2P-Ets129-138), 20 mM NaCl, 2 mM DTT, and ∼10% D2Oat 30 °C and TAZ1 in 20 mM Tris pH 7.0, 50 mM NaCl,2 mM DTT, and ∼10% D2O at 25 °C. The prochiral methyls ofLeu and Val in Ets129-138 were assigned stereospecifically bynonrandom fractional 13C-labeling (14). The spectral assign-ments for Ets129-138 and 2P-Ets129-138 were deposited in the Bio-logical Magnetic Resonance Data Bank under BMRB accessioncodes 4205 and 16426, respectively.

Amide 15N relaxation parameters for 15N-labeled proteinsamples were acquired on a 600-MHz spectrometer at 30 °C,as described previously (15), and analyzed using TENSOR2 (16).

NMR-monitored titrations were carried out by recording sen-sitivity enhanced 15N-HSQC or 13C-HSQC spectra of ∼0.1 mM15N- or 13C∕15N-labeled protein in 20 mM Tris pH 7.0,20–50 mM NaCl as indicated, 2 mM DTT, and ∼10% D2O at25 °C, to which the unlabeled protein partner (∼1.2–2.8 mM stocksolution in the same buffer) was added in small aliquots.

Structural restraints. NOE restraints were determined fromsimultaneous 3D 1H-15N∕13C-1HNOESY-HSQC (heteronuclear

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sequential quantum correlation) (aliphatic side chains),1H-13C-1H NOESY-HSQC (aromatic side chains), and simulta-neous constant-time 3D 1H-15N∕13C-1H NOESY-HSQC spectra(methyl groups), all recorded with τm ¼ 140 ms on a uniformly15N∕13C-labeled protein sample (17).

1HN-15N RDC (residual dipolar coupling) orientation re-straints were obtained from 1H-15N IPAP-HSQC spectra of15N-labeled Ets129-138 and 2P-Ets129-138 recorded in the absencevs. presence of ∼15 mg∕mL and ∼12 mg∕mL Pf1 phage, whichyielded splittings of ∼16.5 Hz and ∼15 Hz in the 2H-NMR spec-tra of the 1HO2H lock solvent, respectively (18). 1Hα-13Cα RDCrestraints were also measured for 2P-Ets129-138 under the sameconditions using an HNCO-based pulse sequence (19, 20).

Ets129-138 structure calculations. Structure calculations were carriedout using Ets129-138 as the spectra of this species are simpler dueto the absence of signals from the disordered residues 1–28 (21).Also, due to rapid hydrogen exchange (HX), helix H0 was notdetected in the initial structure of Ets129-138 determined usingmanually assigned data recorded under higher pH conditionswith a 500-MHz NMR spectrometer (4).

Structure calculations for Ets129-138 were performed usingARIA/CNS v2.1 (22) utilizing distance, dihedral angle, andorientation restraints (Table S1). The majority of the NOESYcross-peaks were manually picked and assigned prior to intensitycalibration by ARIA. Backbone dihedral angles were determinedfrom 13Cα, 13Cβ, 1HN, and 1Hα chemical shifts using the Talos (23)and SSP (secondary structure propensity) algorithms (24). Alimited set of hydrogen bond distance restraints was includedfor amides located in helices that showed protection from rapidexchange after transfer to 2H2O buffer (4). 1HN-15N RDC orien-tation restraints for amides with heteronuclear 1Hf15Ng-NOEratios > 0.2 were then incorporated with the SANI algorithmat iteration 5 of the ARIA protocol, using default energy con-stants of 0.2 and 1 kcalmol−1 Hz−2 for the first and second simu-lated annealing cooling stages, respectively. Values of thealignment tensor (R ¼ 0.6 and Da ¼ þ4.9 Hz) were estimatedby fitting the measured RDC’s against preliminary structuresof Ets129-138 calculated without these restraints. The ARIA/CNS calculations of Ets129-138 were performed as a two-stepprotocol. The initial ARIA round (it0 → it8), starting with anunfolded polypeptide, was completed with only dihedral bondrestraints and uncalibrated NOE data. A second full ARIAcalculation (it0 → it8) was then performed, starting with anunfolded polypeptide and using the final unambiguous andambiguous restraints from the previous round, as well as the

dihedral angle, hydrogen bond, and RDC restraints. A totalof 100 structures per iteration were calculated, with the 25lowest-energy structures of the final iteration refined in a waterbox using Lennard-Jones potentials.

Analyses of the resulting structures were performed withProcheck-NMR (25), Promotif (26), and MolMol (27).

2P-Ets129-138 structure calculations. Structure calculations for2P-Ets129-138 were performed initially using CYANA (28) incor-porating distance, dihedral angle, and orientation restraints, andthen refined using SculptorCNS (29) (Table S1). NOE-baseddistance restraints were generated using the automated NOESYassignment routine in CYANA. Chemical shift derived backbonedihedral angles (with a minimum range of �40°) were includedonly for residues with high TALOS scores (“good > 9”) and inagreement with SSP. Hydrogen bond distance restraints wereincluded only for amides in helices H1–H5 of the core PNTdomain that showed protection from rapid hydrogen exchangein the unmodified protein (4). For subsequent refinement withSculptorCNS, 1HN-15N and 1Hα-13Cα RDC orientationalrestraints for residues with heteronuclear 1Hf15Ng-NOE ratios >0.2 were included in addition to the dihedral angle and CYANA-generated distance restraints. Independent alignment tensorswere used for the dynamic helix H0 and the core PNT domain(H1–H5) as this resulted in lower rms deviations for each of theseregions than obtained with a single tensor or without any RDCrestraints. From total of 500 structures, the 20 lowest-energystructures were selected for the final structural ensemble.

Amide hydrogen exchange. Rapid amide proton-proton exchangerates at 30 °C and pH 6.0 were determined for 15N-labeledEts129-138 and 2P-Ets129-138 by the CLEANEX-PM method(30) using transfer periods ranging from 10 to 60 msec (31). Eachexchange spectrum was recorded using a recycle delay of 1.5 sec,and the reference spectra with a recycle delay of 12.0 sec. Pseudo-first-order rate constants for chemical exchange, kex, wereobtained by least squares fitting of peak intensities using a Matlabmodule provided by J. Choy (University of Toronto, Toronto, ON,Canada) and assuming a negligible contribution from waterrelaxation. HX protection factors (kpred∕kex) were derived usingpredicted kpred values for an unstructured polypeptide with thesequence of Ets129-138, calculated with the SPHERE server(32, 33) using poly-D,L-alanine reference data corrected foramino acid type, pH, temperature, and isotope effects, andasparate as surrogate for phosphoserine/threonine.

1. Foulds CE, Nelson ML, Blaszczak AG, Graves BJ (2004) Ras/mitogen-activated proteinkinase signaling activates Ets-1 and Ets-2 by CBP/p300 recruitment. Mol Cell Biol24:10954–10964.

2. Seidel JJ, Graves BJ (2002) An ERK2 docking site in the Pointed domain distinguishes asubset of ETS transcription factors. Genes Dev 16:127–137.

3. Gunther CV, Graves BJ (1994) Identification of ETS domain proteins in murine Tlymphocytes that interact with the Moloney murine leukemia virus enhancer.Mol CellBiol 14:7569–7580.

4. Slupsky CM, et al. (1998) Structure of the Ets-1 pointed domain and mitogen-activatedprotein kinase phosphorylation site. Proc Natl Acad Sci USA 95:12129–12134.

5. Mackereth CD, et al. (2004) Diversity in structure and function of the Ets family PNTdomains. J Mol Biol 342:1249–1264.

6. McIntosh LP, et al. (2009) Detection and assignment of phosphoserine and phospho-threonine residues by 13C-31P spin-echo difference NMR spectroscopy. J Biomol NMR43:31–37.

7. Kang HS, et al. (2008) Identification and structural characterization of a CBP/p300-binding domain from the ETS family transcription factor GABP alpha. J Mol Biol377:636–646.

8. Waas WF & Dalby KN (2002) Transient protein-protein interactions and a random-or-dered kinetic mechanism for the phosphorylation of a transcription factor by extra-cellular-regulated protein kinase 2. J Biol Chem 277:12532–12540.

9. Fredricksen RS, Swenson CA (1996) Relationship between stability and function forisolated domains of troponin C. Biochemistry 35:14012–14026.

10. Delaglio F, et al. (1995) NMRPipe: A multidimensional spectral processing system basedon UNIX pipes. J Biomol NMR 6:277–293.

11. Goddard TD, Kneller, DG. Sparky 3. (Univ of California, San Francisco).

12. Sattler M, Schleucher J, Griesinger C (1999) Heteronuclear multidimensional NMRexperiments for the structure determination of proteins in solution employing pulsedfield gradients. Prog NMR Spect 34:93–158.

13. Slupsky CM, Gentile LN, McIntosh LP (1998) Assigning the NMR spectra of aromaticamino acids in proteins: Analysis of two Ets pointed domains. Biochem Cell Biol76:379–390.

14. Szyperski T, Neri D, Leiting B, Otting G, Wuthrich K (1992) Support of 1H NMR assign-ments in proteins by biosynthetically directed fractional 13C-labeling. J Biomol NMR2:323–334.

15. Farrow NA, Zhang O, Forman-Kay JD, Kay LE (1994) A heteronuclear correlationexperiment for simultaneous determination of 15N longitudinal decay and chemicalexchange rates of systems in slow equilibrium. J Biomol NMR 4:727–734.

16. Dosset P, Hus JC, Blackledge M, Marion D (2000) Efficient analysis of macromolecularrotational diffusion from heteronuclear relaxation data. J Biomol NMR 16:23–28.

17. Zwahlen C, et al. (1998) An NMR experiment for measuring methyl-methyl NOEs inC-13-labeled proteins with high resolution. J Am Chem Soc 120:7617–7625.

18. Ottiger M, Delaglio F, Bax A (1998) Measurement of J and dipolar couplings fromsimplified two-dimensional NMR spectra. J Magn Reson 131:373–378.

19. Yang DW, Tolman JR, Goto NK, Kay LE (1998) An HNCO-based pulse scheme for themeasurement of C-13(alpha)-H-1(alpha) one-bond dipolar couplings in N-15, C-13labeled proteins. J Biomol NMR 12:325–332.

20. de Alba E, Tjandra N (2004) Residual dipolar couplings in protein structure determina-tion. Methods Mol Biol 278:89–106.

21. Macauley MS, et al. (2006) Beads-on-a-string, characterization of ETS-1 sumoylatedwithin its flexible N-terminal sequence. J Biol Chem 281:4164–4172.

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22. Rieping W, et al. (2007) ARIA2: Automated NOE assignment and data integration inNMR structure calculation. Bioinformatics 23:381–382.

23. Cornilescu G, Delaglio F, Bax A (1999) Protein backbone angle restraints from search-ing a database for chemical shift and sequence homology. J Biomol NMR 13:289–302.

24. Marsh JA, Singh VK, Jia Z, Forman-Kay JD (2006) Sensitivity of secondary structurepropensities to sequence differences between alpha- and gamma-synuclein: implica-tions for fibrillation. Protein Sci 15:2795–2804.

25. Laskowski RA, Rullmannn JA, MacArthur MW, Kaptein R, Thornton JM (1996) AQUAand PROCHECK-NMR: Programs for checking the quality of protein structures solvedby NMR. J Biomol NMR 8:477–486.

26. Hutchinson EG, Thornton JM (1996) PROMOTIF—a program to identify and analyzestructural motifs in proteins. Protein Sci 5:212–220.

27. Koradi R, Billeter M, Wuthrich K (1996) MOLMOL: A program for display and analysisof macromolecular structures. J Mol Graph 14:51–55, 29–32.

28. Guntert P (2004) Automated NMR structure calculation with CYANA. Methods MolBiol 278:353–378.

29. Charavay C, Eynard J, Hus J, Bouvignies G, Blackledge M. SculptorCNS:Biomolecular structure determination and refinement in solution from NMR residualdipolar couplings, paramagnetic phenomena and spin relaxation data.

30. Hwang TL, van Zijl PC, Mori S (1998) Accurate quantitation of water-amide protonexchange rates using the phase-modulated CLEAN chemical EXchange (CLEANEX-PM) approach with a Fast-HSQC (FHSQC) detection scheme. J Biomol NMR 11:221–226.

31. Lee GM, et al. (2005) The structural and dynamic basis of Ets-1 DNA bindingautoinhibition. J Biol Chem 280:7088–7099.

32. Bai Y, Milne JS, Mayne L, Englander SW (1993) Primary structure effects on peptidegroup hydrogen exchange. Proteins 17:75–86.

33. Zhang YZ (1995) Protein and peptide structure and interactions studied by hydrogenexchange and NMR. PhD thesis (Univ of Pennsylvania, Philadelphia).

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Fig. S1. CBP TAZ1 domain binds Ets1 and Ets2 PNT domains. (A) Schematic of CBP with amino acid coordinates and known domains. (B) FLAG-HMK-Ets11-440

(full length), His6-FLAG-HMK-Ets11-138, His6-FLAG-HMK-Ets21-172, were expressed, purified, and ERK2 phosphorylated in vitro or mock-treated, as previouslydescribed (1, 2). GST-tagged CBP species were expressed in Rosetta cells (Novagen) and purified on GSTrap FF columns (GE Healthcare). FLAG pull-downexperiments were conducted as previously described (1), except that NP-40 was omitted from all reaction buffers. Briefly, 240 pmol of the FLAG-taggedEts1 and Ets2 species were captured on FLAG antibody beads (Sigma) and GST-CBP fragments were added to binding reaction at 400 nM for 4 hr at 4 °C.Proteins (bound GST-CBP and preloaded FLAG-Ets1/2 species) were eluted from the FLAG beads with SDS-sample buffer, resolved by SDS-PAGE gels anddetected by immunoblotting with α-GST (GE Healthcare) for GST-CBP fusions, α-His (His Probe Santa Cruz) for His6-FLAG-HMK-Ets11-138,His6-FLAG-HMK-Ets21-172, or α-Ets1 (UT2) antibodies (3) for FLAG-HMK-Ets1. Lane 1 shows 4% of the GST-CBP input proteins tested for binding. Lanes2–8 show GST-CBP input proteins bound to beads or to the indicated Ets1/2 species. The loading controls display an aliquot of loaded beads to confirm thatan equal amount of each FLAG-tagged protein was loaded on FLAG antibody beads. (Molecular masses were as expected for tagged species, but not indicated.)Phosphorylation of Ets1 and Ets2 is apparent by the slight decrease in mobility on the immunoblot. The positions of molecular mass markers (kDa) for GST-CBPfragments are indicated. Although some immunoblots showmodest effects of phosphorylation of Ets1 and Ets2 species on CBP binding, these differences wereimpossible to quantify with this assay. ITC experiments were used to measure a ∼34-fold effect of phosphorylation on the affinity of the isolated TAZ1 domainfor Ets11-138.

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A B

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Fig. S2. ETS family PNT domain structural elements and sequence conservation. (A) Dendogram of selected family members based on their PNT domainsequences using ClustalW (1). (B) Helical elements of the PNT domain. ERG (PDB ID code 1sxe), Tel (PDB ID code 1lky), Yan (PDB ID code 1sv4), and Fli1(PDB ID code 2ytu) share a core helical bundle (H2–H5) also common to SAM domains, whereas GABPα (PDB ID code 1sxd) and SPDEF (PDB ID code2dkx) have an N-terminal helix H1 (5). Those phosphorylated byMAPK have additional observed (Ets1, Ets2) or predicted (Pnt-P2) helices H0 and H1. (C) Portionsof the murine Ets1, Ets2, GABPα, Tel, and Fli1, human SPDEF and ERG, and Drosophila Pnt-P2 and Yan sequences aligned by ClustalW. Residues with similarphysicochemical properties as those in Ets1, Ets2, or Pnt-P2 are shaded gray, and Ets1 phosphoacceptors Thr38 and Ser41 aremarked by asterisks. Numbers referto amino acid positions in the full-length proteins and to the helical boundaries in Ets1. (D) SSP scores (24) derived from 15N, 1HN, 13Cα, and 13Cβ chemical shiftsconfirm that Ets269-172 has the same helical secondary structure as Ets129-138 and that helix H0 is preceded by unstructured phosphoacceptors Thr72 andpotentially Ser75. The helices forming the PNT domain have scores approaching 1, whereas the unstructured N-terminal region has scores near 0. Missingdata corresponds to prolines or amides lacking unambiguous spectral assignments. The NMR spectra of Ets263-172 in 20 mM phosphate pH 7.0, 20 mM KCl,0.1 mM EDTA, and 2 mM DTT, 25 °C were assigned by standard methods.

1. Larkin MA, et al. (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948.

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Fig. S3. Helix H0 and the phosphoacceptors Thr38 and S41 are dynamic in both Ets129-138 (red) and 2P-Ets129-138 (blue). However, phosphorylation leads toslight stabilization of helix H0 against local unfolding and a small increase in average helical propensity at the H0/H1 junction. (A) SSP scores, calculated frommainchain 1H, 13C, and 15N chemical shifts, increase from 0 to þ1 with increasing α-helical propensity (24). (B–D) 15N T1, T2, and 1Hf15Ng-NOE relaxation dataand errors, recorded at 30 °C with a 600-MHz NMR spectrometer. (E) Generalized order parameters, S2, from an anisotropic model free analysis of 15N relaxa-tion data using TENSOR2 (16). To extract the S2 values describing internal mobility, low-energy members of the structural ensembles of Ets129-138 and2P-Ets129-138 were first used to determine their diffusion tensors for global rotation. These tensors corresponded to isotropic tumbling times of7.1� 0.04 nsec and 6.3� 0.02 nsec for the two monomeric species, respectively. Note that reduced NOE and S2 values and increased T2 values reflect increasedmobility of the amide 15N-1HN bond vector on a nsec-psec time scale. (F) Protection factors (PFs) derived from the ratio of predicted vs. observed rates of amideHX. The low PFs of residues in helix H0 demonstrate that this helix is marginally stable and undergoes facile local unfolding and exchange. (G) Relative protec-tion factors for 2P-Ets129-138 vs. Ets129-138. The modest increase in corresponding PFs indicates that phosphorylation slightly stabilizes helix H0 against unfold-ing. Missing data correspond to prolines, residues before prolines, residues with unresolved 1HN-15N signals, or in the case ofD and E, residues that do not showmeasurable HX (pH 6.0, 30 °C) by the CLEANEX method and thus have protection factors > 10. The phosphoacceptors Thr38 and Ser41 are indicated witharrows. The dynamic nature of helix H0 is not a trivial result of deleting the disordered residues 1–28, as the NMR spectra of corresponding residues inEts11-138 overlap those of Ets129-138 closely, confirming identical features (21).

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L49 Cδ2

A48 Cβ

Fig. S4. Helix H0 of Ets11-138 is conformationally dynamic and sensitive to trypsin proteolysis. Phosphorylation of Ets11-138 promoted local protease resistanceand changes in CD spectra, but not global thermal stability. (A) Schematic of Ets11-138 helical secondary structure (colored boxes) and potential trypsin cleavagesites (ticks). The gray box denotes the N-terminal His6-FLAG-HMK-tag. Mass spectrometry identified cleavage sites are indicated by amino acid position (2, 15,42, and 50 result from lysine-directed cleavages, whereas 62 is an arginine cleavage). (B) SDS-PAGE gels of partial digests of unmodified Ets11-138 (Upper) and2P-Ets11-138 (Lower) as a function of increasing trypsin concentration. The preferential cleavage at Lys42, Lys50, and Arg62 within helices H0 and H1 revealed atleast transient accessibility of these residues to trypsin, whereas lysines and arginines within the core PNT domain were more resistant to proteolysis. Thetransient accessibility of residues in helices H0 and H1 was reduced upon phosphorylation, indicative of local stabilization. Cleaved fragments are denotedby position of N-terminal cleavage site, and all fragments terminated at position 138. ♦, position of trypsin. (C) Thermal unfolding curves of Ets11-138

(Tm ¼ 77� 1 °C) and 2P-Ets11-138 (76� 1 °C), as monitored by CD. The global stability of Ets11-138 against thermal unfolding did not change significantly uponphosphorylation. This is consistent with the NMR-based observations that Ets151-138, which lacks helix H0, still adopts a stably folded structure, whereas Ets11-52

is unstructured. However, the mean residue molar ellipticity ½θ�222 of Ets11-138 decreased by ∼7% from −6;500 to −7;000 deg cm2∕dmol upon dual phosphor-ylation of Thr38 and Ser41. This is suggestive of slightly increased helical secondary structure content and/or perturbations in relative helix orientations (35). (D)Loss of long-range 1H-1H NOEs demonstrates that phosphorylation leads to displacement of helix H0 from the core PNT domain. Shown are strip plots fromconstant-timemethyl 13C-13C-1HNOESY spectra of (left strip of pair) Ets129-138 and (right strip) 2P-Ets129-138 at the 13Cmethyl shifts of Leu49 and Ile124 (17). KeyNOEs that defined the position of helix H0 in the unmodified protein are identified in red boxes. These are uniformly absent in 2P-Ets129-138.

1. Gagne SM, et al. (1994) Quantification of the calcium-induced secondary structural changes in the regulatory domain of troponin-C. Protein Sci 3:1961–1974.

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11 10 9 8 7 6

130

125

120

115

110

H0 H1 H2 H2’ H3 H4 H5PP

Ets

129-

138

Ets

142-

138

Ets

151-

138

9.4 9.2 9.0 8.8

111

110

109

108

15N

(p

pm

)

G55

G102H128 L125

Ets

11-1

38

1H (ppm)

8.0 7.8 7.6 7.4

120

119

118

117

A

B

C2P

-Ets

11-1

38

*

Fig. S5. Colinear amide chemical shift changes in the Ets1 PNT domain upon deletion or phosphorylation supports the hypothesis that helix H0 exists in aequilibrium between open and closed states. (A) N-terminal deletion mutants of Ets1. Arrows denotes the starting residue of the construct or the sites ofphosphorylation. (B) Overlaid 15N-HSQC spectra of the deletion mutants and of phosphorylated 2P-Ets11-138, color coded according to A. An aliased signal,displaced in the 15N dimension due to use of different spectral widths, is indicated by an *. (C) Colinear amide chemical shift changes are observed for some H0/H2/H5 interfacial residues, such as Gly55 (Left), and His128 and Leu125 (Right). Leu125 and His128, which are in the core PNT domain, have similar amide1HN-15N chemical shifts and hence structural roles in three nonphosphorylated fragments containing H0 (Ets11-138, Ets129-138, and Ets142-138). However, asexpected due to the packing of H0 against these core PNT domain residues, deletion of this helix (Ets151-138) leads to chemical shift changes for Leu125and His128. Importantly, in phosphorylated 2P-Ets11-138, the amide chemical shifts of Leu125 and His128 are intermediate between those in the unmodifiedproteins with vs. without helix H0. The simplest interpretation of these colinear chemical shift changes is that phosphorylation perturbs a conformationalequilibrium between a closed state with helix H0 packed against the core PNT domain and an open state with H0 fully displaced (or deleted). This occurs in thefast exchange limit, yielding population-weighted chemical shifts. Furthering this argument, the amide of Gly55, located at the H0/H1 junction, appears to be avery sensitive indicator of the conformation of the PNT domain. Based on the structures of the Ets1 fragments, we propose that Gly55 partakes in hydrogenbonding interactions that depend upon the continuity (i.e., bent vs. linear; see Fig. 2) of the two helices flanking this junction. In particular, more downfield 1HN

shifts, which result from stronger hydrogen bonding, likely reflect H0 being displaced from the PNT domain and more continuous with H1. As with Leu125 andHis128, Gly55 reports colinear amide chemical shift changes, moving downfield in the series Ets11-138ð¼ Ets129-138Þ to 2P-Ets11-138 to Ets151-138. However, inEts142-138 (deleted to the start of H0), the amide of Gly55 still follows this colinear pattern but shifts further upfield from its spectral position in Ets151-138. Inother words, deletion of residues 1–41 appears to allow H0 to adopt an even more closed conformation (i.e., with the H0/H1 junction bent) than in the longerunmodified Ets1 fragments. Alongwith several lines of evidence that helix H0 is dynamic, these simple spectral comparisons strongly support ourmodel that H0exists in a conformational equilibrium between closed and open states in unmodified Ets1 and that phosphorylation shifts the population distribution towardthe open state.

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9.0 8.5 8.0 7.59.51H (ppm)

130

125

120

115

110

15N

(p

pm

)

G55G92

F120

T119

K91

G122

G109

S89

L125

S54

S40

F53S41

L37

A51

L129

T52

M44

L49

K50

D123

L36

Q47

F53

C112

L129

M44

L36N45

V93

G55G92

S41T38

W126

D94G109

G122R32

A3

A26

K91D119

V81F120

S40

L132

S54

T52

L125

S89

V34E43

V85M82

D20

Q60

Q133

F88

S46

D33K50

D123

L49

H128

I124

N86L90

9.0 8.5 8.0 7.59.5

Ets11-138 2P-Ets11-138A Amides

8.5 7.5 6.5 5.5

135

130

125

120

115

MethylsC

9.0 8.5 8.0 7.59.5

130

125

120

115

11015

N (

pp

m)

13C

(p

pm

)

Salt effectB

2P-Ets11-138:TAZ1(1:0, 1:0.5, 1:1)

Ets11-138:TAZ1(1:0, 1:0.5, 1:1)

Ets11-138

G55

G92

F120

S40

T52

L125

S89

G122

G109

F53 L129

D119K91

A51

L37

L36

Q47

L49K50

D123

no TAZ1(20mM NaCl)

1:1 TAZ1(500mM NaCl)

S41

S54

W80ζ2W72ζ2

W72ε3

W83δ1

H128ε1

F88ε*F120ε*

W83η2

F88δ*

Ets129-138:TAZ1(1:0, 1:1)

W72ζ3

W72δ1W72η2

W80δ1

2.5 2.0 1.5 1.0 0.5 0.030

25

20

15

D Aromatics

1H (ppm)

13C

(p

pm

)

T52γ2*

L129δ1*

L129δ2*

L125δ2*

I124γ2*

I124δ1*

V121γ2*

A117β*

L105δ1*

A51β*

L49δ2*

A48β*

Ets129-138:TAZ1(1:0, 1:1, 1:4)

Ets129-138 Ets129-138

H (ppm)

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0

0.2

0.4

0.6

0.8

1.0

inte

nsi

ty r

atio

1 11 21 31 41 51 61 71 81 91 101 111 121 131

1 11 21 31 41 51 61 71 81 91 101 111 121 131residue

0

0.2

0.4

0.6

0.8

1.0

inte

nsi

ty r

atio

T38

S41

H1H0 H2 H4 H5H2’ H3E

F

H0H0

H5H2

H5H0

H2

H5

H0

H2

H5

H1

H1

H1

H1

H4

H3

H2’

H2’ H4

H3

H2’

Ets11-138 2P-Ets11-138

pT38

pS41

pT38

pS41T38

S41T38

S41

Ets11-138:TAZ1(1:0 vs 1:0.5, 1:0 vs 1:1)

2P-Ets11-138:TAZ1(1:0 vs 1:0.5, 1:0 vs 1:1)

Fig. S6. The TAZ1-binding interface on Ets11-138 is localized primarily to helices H0 and H5. (A) Overlaid 15N-HSQC spectra of ∼0.1 mM 15N-labeled (Left)Ets11-138 and (Right) 2P-Ets11-138 titrated with unlabeled TAZ1 in molar ratios of 1∶0 (light blue), 1∶0.5 (dark blue), and 1∶1 (red), recorded for samples in20 mM Tris pH 7.0, 20 mM NaCl. Based on the protein concentrations and the Kd values from ITC measurements, these ratios correspond to 0, ∼30%,and ∼50% saturation of Ets11-138 and 0, ∼50%, and ∼90% saturation of 2P-Ets11-138, respectively. Amides showing >85% loss of signal intensity uponTAZ1 binding are labeled. (B) Increased ionic strength disrupts binding as evidenced by the reappearance of many amide signals from 15N-labeled (Left)Ets11-138 in the presence of 1∶1 TAZ1 upon addition of NaCl to 500 mM. The labeled peaks are significantly weaker when the complex is studied underthe lower ionic strength conditions of A. Note that, although some amides show small ionic strength-dependent chemical shift changes, the folded structureof Ets11-138 is still retained at 500mMNaCl. (C andD) Overlaid 13C-HSQC spectra of the methyl and aromatic signals of Ets129-138 titrated with unlabeled TAZ1 inmolar ratios of 1∶0 (light blue), 1∶1 (red; ∼40% saturation), and 1∶4 (black;∼80% saturation), recorded for samples in 20mMTris pH 7.0, 50mMNaCl. The peaksshowing the most pronounced intensity changes upon TAZ1 binding are labeled. (E) Histograms of relative amide signal intensity at the indicated equivalenceratios relative to unbound Ets11-138 (Top) and 2P-Ets11-138 (Bottom). (F) Residues with >0.85 loss of amide signal intensity upon titration to a 1∶1 ratio aremapped onto corresponding low-energy members of the structural ensembles of (Left) Ets129-138 and (Right) 2P-Ets129-128 Increasing cyan sphere size corre-sponds to greater signal loss upon complex formation (intensity ratios: 0 < large sphere < 0.05, 0.05 < medium < 0.10, 0.10 < small < 0.15). Missing data pointscorrespond to prolines or residues with unassigned or overlapping NMR signals. It is possible that conformational exchange between free and bound states orwithin the bound state occurring on an unfavorable time scale relative to the chemical shift differences between these states leads to severe linewidth broad-ening and, therefore, the loss of detectable signals, particularly in context of the increased molecular mass of the complex. The latter scenario is favored assignals from the bound complex are not observed at the 80 to 90% saturation reported here.

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0

0.2

0.4

0.6

0.8

1.0

inte

nsi

ty r

atio

340 350 360 370 380 390 400 410 420 430residues

0

0.2

0.4

0.6

0.8

1.0

inte

nsi

ty r

atio

H1 H2 H4H3

A

B

C

H1

TAZ1+Ets11-138 TAZ1+2P-Ets11-138

H2

H3

H4 H1

H2

H3

H4H1

H2

H3

H4H1

H2

H3

H4

1H (ppm)

15N

(p

pm

)

9.0 8.5 8.0 7.5 7.0

125

120

115

110

Q355

I414

C429

9.0 8.5 8.0 7.5 7.0

K365N434

S410S416

H417

I414V428Q367

C429

H362

Q354W418

A372

TAZ1 TAZ1

340 350 360 370 380 390 400 410 420 430

residues

K419H364

K351

R368

H362

TAZ1:Ets11-138

(1:0 vs 1:1)

TAZ1:2P-Ets11-138

(1:0 vs 1:0.66, 1:0 vs 1:1)

TAZ1:Ets11-138

(1:0, 1:1)TAZ1:2P-Ets11-138

(1:0, 1:0.66, 1:1)

Fig. S7. The Ets1-binding interface on TAZ1 localizes to an extended interface. (A) Overlaid 15N-HSQC spectra of ∼0.1 mM 15N-labeled TAZ1 titrated with (Left)Ets11-138 and (Right) 2P-Ets11-138 in molar ratios of 1∶0 (light blue), 1∶0.5 (dark blue), and 1∶1 (red), recorded for samples in 20 mM Tris pH 7.0, and 20 or 50 mMNaCl, respectively. Based on the protein concentrations and the ITC-determined Kd values, these ratios correspond to 0 and ∼50% saturation of TAZ1 withEts11-138 and 0, ∼60%, and ∼90% saturation of with 2P-Ets11-138, respectively. Amides showing >95% loss of signal intensity upon TAZ1 binding are labeled.(B) Histograms of TAZ1 amide signal intensity at the indicated equivalence ratios. Missing data points correspond to prolines or residues with unassigned oroverlapping NMR signals. (C) Residues showing >0.9 loss of signal intensity upon titration to a 1∶1 ratio with (Left) Ets11-138 and (Right) 2P-Ets11-138 are mappedonto a low-energy member of the structural ensembles of TAZ1. Increasing cyan sphere size corresponds to greater signal loss upon complex formation(intensity ratios: 0 < large sphere < 0.05, 0.05 < small < 0.10). The Znþ2 ions of TAZ1 are shown as magenta balls.

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130

120

110

100

80

60

40

20

0

inte

nsi

ty lo

ss (

%)

15N

(p

pm

)

9.5 8.5 7.5

PP

29-1

38

42-1

38

51-1

38

1-13

8

2P 1

-138

H0 H1 H2 H3 H4 H5H2’• ••

A B C

D

F

1H (ppm)

9.5 8.5 7.5 9.5 8.5 7.5

9.4 9.3 9.2 9.1 9.0 8.9

111

110

109

108

1H (ppm)

15N

(p

pm

)

EGly102

Gly55

Ets1 29-138-L49R Ets129-138 -D123R Ets1 29-138-E127R

29-1

38

42-1

38

51-1

38

2P 1-

138

L49R

D123R

E127R

L49

D12

3E

127

open

closed

Gly55 Gly55Gly55

WT

T52

A

L49

E(1

)

T38

A/S

41A

S41

A

T38

A

T52

R

F53

A

D11

9R(1

)

αpT38–Ets1

αEts1

K50

A(2

)E

127R

D11

9A

W12

6A

E12

7A(2

)

WT

E12

7A(1

)

E12

7R

WT

D11

9A

W12

6A

E12

7A(2

)

TW

) 2(E94

L

) 1(R94

L

R321D

) 2(R911

D

R321/ 911D

) 2(A45

S

A321D

TW

) 2(A911

D

) 3(R911

D

A321D

) 2(R321

D

R321D

R321/ 911D E

127A

(1)

WT

K52

A(2

)

G

R34E

αEts1

WT

)3(E94

L

) 2(R94

L

WT

S54

A(1

)

K50

A

WT

αpT38–Ets1

Fig. S8. Mutation of the TAZ recognition interface of the Ets1 PNT domain disrupts TAZ1 binding without perturbing global folding, expression, or phos-phorylation. (A–C) 15N-HSQC spectra of 15N-labeled Ets129-138 with the point mutations L49R (cyan) in helix H0 and D123R (dark orange) or E127R (light orange)in helix H5. In each case, the protein remains stably folded, giving well-dispersed spectra. This demonstrates that disrupted binding and reduced in vivo func-tion are not an indirect consequence of the mutations perturbing the global fold of the PNT domain. (D) A small portion of the overlaid spectra of these threemutants, along with various deletion fragments and phosphorylated proteins, color coded according to the cartoon of panel (E). The amide chemical shifts ofGly55 in these three mutants are colinear with the species discussed previously in Fig. 2D and Fig. S5. This indicates that the mutations, which lie along theinterface between helices H0 and H1, all shift the population distribution of the PNT domain toward the open state. The small deviation of the amide signal ofGly55 in Ets151-138 (red) from this exact colinear pattern likely results from the presence of the newly introduced N terminus at position 51. (F) Although shiftingthe PNT domain toward the open state, each of the three mutants weakens TAZ1 binding, presumably due to the introduction of an unfavorable positivecharge at the complex interface. The weakening of binding is shown by the average reduced intensity of four amides (Gly55, Ser89, Gly122, and Leu125) upontitration to a 1∶1 molar ratio with labeled TAZ1. Note that the intensity losses for 2P-Ets11-138 (magenta), Ets129-138 (green), and Ets142-138 (blue) reflect therelative affinities of these species for TAZ1. In contrast, Ets152-138 (red) does not measurably bind TAZ1 due to the deletion of all residues forming helix H0. (G)Controls for mutant Ets1 species presented in Figs. 1 and 4. For expression controls and phosphorylation status, ∼1.5 × 106 cells were transfected with 3 μg offull-length FLAG-Ets1 expression plasmid, 3 μg constitutively active MEK1, and 50 mL lipofectamine (Invitrogen). As described previously (1), antibodies againstthe FLAG tag were used for immunoprecipitation, followed by immunoblot detection with either anti-Ets1 antisera (UT2) (3) or phospho-specific antiseraagainst pThr38-Ets1 (Biosource). All experiments had WT controls performed in parallel with assay of mutants, as shown. Full-length Ets1 species have ex-pression levels similar to WT. Phosphorylation of Thr38 is also very similar among the mutants except for mutants with a phosphoacceptor site substitution(T38A, T38A;S41A, and T38E;S41E). The (#) designation refers to version of replica data within this figure. Upper rows show immunoblots from assays in whichexpression (Top) and Thr38 phosphorylation (Bottom) status was determined in the same transfected cell extracts. The middle row shows immunoblots fromassays in which only expression was determined. The bottom row shows immunoblots from assays in which only Thr38 phosphorylation status was determined.Two methods were used for secondary antibody system for pThr38-Ets1 detection: ECL Plus (Amersham) with horseradish peroxidase (darker panels, also inupper immunoblot), and ECL Plex (Amersham) with Cy5 (lighter panels).

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Table S1. NMR restraints and structural statistics for Ets129-138 and 2P-Ets129-138 ensembles

Ets129-138 2P-Ets129-138

Summary of restraintsNOEs *

Intraresidue 781 (151) 511Sequential 572 (119) 510Medium range (1 < ji − jj < 5) 763 (251) 498Long range (ji − jj ≥ 5) 693 (302) 529Total 2,809 (823) 2,048

Dihedral anglesϕ, ψ , χ1 89, 89, 0 91, 91, 0

Hydrogen bonds 17 × 2 24 × 2Residual dipolar couplings 91(15N-1HN) 81 (15N-1HN), 71 (13Cα-1Hα)Residues in allowed regions of Ramachandran plot † 99.70% 99.50%

Mean energies, kcal-mol−1

Evdw −104.7 ± 21.2 −571.9 ± 13.2Ebonds 51.2 ± 2.3 18.6 ± 0.6Eangles 263.8 ± 12.3 73.9 ± 2.7Eimpr 167.6 ± 11.4 15.1 ± 1.0ENOE 347.9 ± 16.4 120.9 ± 9.6Ecdih 20.3 ± 3.7 0.5 ± 0.2Esani 142.2 ± 6.2 133.6 ± 3.3

rms deviation, ÅFor structured elements ‡ including H0

Backbone (N, Cα, C’) 0.23 1.81All heavy atoms 0.67 2.19

For structured elements ‡ excluding H0Backbone (N, Cα, C’) 0.21 0.29All heavy atoms 0.67 0.88

*Ambiguous NOEs are shown in parenthesis for Ets129-138.†Calculated with Procheck-NMR (25), and summed over most favored, allowed, and generously allowedregions.

‡Consensus secondary structure, calculated from Promotif (25). H0, 42-52; H1, 54-62; H2, 74-87; H2′, 95–98(310 helix); H3, 102–107; H4, 109–116; H5, 119–134.

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