Characterizing Watson–Crick versus Hoogsteen Base Pairing in a

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Characterizing Watson−Crick versus Hoogsteen Base Pairing in a DNA−Protein Complex Using Nuclear Magnetic Resonance and SiteSpecifically 13C- and 15N‑Labeled DNA Huiqing Zhou,†,@ Bharathwaj Sathyamoorthy,‡ Allison Stelling,† Yu Xu,§ Yi Xue,∥ Ying Zhang Pigli,⊥ David A. Case,# Phoebe A. Rice,⊥ and Hashim M. Al-Hashimi*,†,§ Biochemistry Downloaded from pubs.acs.org by UNIV OF LOUISIANA AT LAFAYETTE on 04/06/19. For personal use only.

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Department of Biochemistry, Duke University School of Medicine, Durham, North Carolina 27710, United States Department of Chemistry, Indian Institute of Science Education and Research Bhopal, Bhopal 462066, India § Department of Chemistry, Duke University, Durham, North Carolina 27708, United States ∥ Tsinghua-Peking Center for Life Sciences, School of Life Sciences, Tsinghua University, Beijing 100084, China ⊥ Biochemistry and Molecular Biology, The University of Chicago, Chicago, Illinois 60637, United States # Department of Chemistry and Chemical Biology, Rutgers University, Piscataway, New Jersey 08854, United States ‡

S Supporting Information *

ABSTRACT: A(syn)-T and G(syn)-C+ Hoogsteen base pairs in protein-bound DNA duplexes can be difficult to resolve by X-ray crystallography due to ambiguous electron density and by nuclear magnetic resonance (NMR) spectroscopy due to poor chemical shift dispersion and size limitations with solution-state NMR spectroscopy. Here we describe an NMR strategy for characterizing Hoogsteen base pairs in protein−DNA complexes, which relies on site-specifically incorporating 13 C- and 15N-labeled nucleotides into DNA duplexes for unambiguous resonance assignment and to improve spectral resolution. The approach was used to resolve the conformation of an A-T base pair in a crystal structure of an ∼43 kDa complex between a 34 bp duplex DNA and the integration host factor (IHF) protein. In the crystal structure (Protein Data Bank entry 1IHF), this base pair adopts an unusual Hoogsteen conformation with a distorted sugar backbone that is accommodated by a nearby nick used to aid in crystallization. The NMR chemical shifts and interproton nuclear Overhauser effects indicate that this base pair predominantly adopts a Watson−Crick conformation in the intact DNA−IHF complex under solution conditions. Consistent with these NMR findings, substitution of 7-deazaadenine at this base pair resulted in only a small (∼2-fold) decrease in the IHF−DNA binding affinity. The NMR strategy provides a new approach for resolving crystallographic ambiguity and more generally for studying the structure and dynamics of protein−DNA complexes in solution.

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ollowing the discovery of the DNA double helix,1 much effort was directed toward determining single-crystal X-ray structures of isolated purine-pyrimidine dimers to test the fine details of Watson−Crick base pairing proposed by Watson and Crick.2−8 The first successful attempt was reported in 1959, when Karst Hoogsteen determined the X-ray co-crystal structure of base pair derivatives containing 9-methyladenine and 1-methylthymine.2 Rather than an A-T Watson−Crick base pair (bp), an alternative pairing mode called “Hoogsteen” was observed (Figure 1A). Relative to a Watson−Crick bp, the adenine base flips 180° to adopt a syn rather than an anti conformation (Figure 1A). The base forms a unique set of hydrogen bonds (H-bonds) that require that the two bases come into closer proximity by ∼2.0−2.5 Å3,9 (Figure 1A), which in turn induces a kink in the DNA double helix.10,11 Analogous G-C+ Hoogsteen bps12 form by flipping the guanine base but also require protonation of cytosine N3 to form a second H-bond (Figure 1A). © XXXX American Chemical Society

Since their discovery, Hoogsteen bps have been observed in X-ray structures of AT-rich sequences that form duplexes entirely composed of Hoogsteen bps;14−17 in structures of DNA bound to small molecule bis-intercalators;12,18−22 in damaged DNA bases incapable of forming Watson−Crick bps where they are thought to play roles in damage accommodation,23−25 recognition,26 and repair;27 and in Y-family “low-fidelity polymerases” that replicate DNA using Hoogsteen pairing as a means of bypassing lesions on the Watson−Crick face during replication.28−31 More recently, nuclear magnetic resonance (NMR) relaxation dispersion (RD) methods32−34 revealed that in canonical duplex DNA, A-T and G-C Watson−Crick bps exist in dynamic equilibrium with shortlived (lifetimes of 0.2−2.5 ms) and sparsely populated (0.1− 5%) Hoogsteen bps.11,35−39 Received: January 11, 2019 Revised: February 28, 2019

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Figure 1. Structures of Watson−Crick and Hoogsteen base pairs and crystal structure of the IHF−DNA complex. (A) Chemical structures of Watson−Crick and Hoogsteen base pairs. Differences in hydrogen bonding, cytosine protonation, purine base orientation (anti vs syn), and C1′− C1′ distance are highlighted. Short atomic distances involving imino protons that give rise to characteristic nuclear Overhauser effects in A-T Watson−Crick (AH2-TH3) and Hoogsteen (AH8-TH3) bps are indicated using black arrows. (B) X-ray structure of the IHF−DNA complex (Protein Data Bank entry 1IHF13). The consensus sequence for specific protein recognition is colored blue. The A21-T48 Hoogsteen-like (HGlike) bp, the A56-T13 Watson−Crick (WC) bp, and the control A9-T60 Watson−Crick bp are shown in red, orange, and gray rectangles, respectively. The nick in one DNA strand is indicated with an arrow. Two pseudosymmetry-related large kink sites are marked with dashed lines in the DNA sequence, corresponding to positions of intercalating proline residues.

assign a Watson−Crick versus Hoogsteen geometry. Subsequent structural and biochemical studies, including studies examining the impact of single-atom substitutions that destabilize Hoogsteen pairing in the DNA template, confirmed that this polymerase does indeed employ a Hoogsteen mechanism to replicate certain base pairs.30,48 Other studies have noted the difficulty in resolving Hoogsteen versus Watson−Crick bps in X-ray structures of protein−DNA complexes, including the DNA homeodomain45 and DNA− p73 complexes.46 Thus, it is conceivable that other Hoogsteen bps in crystal structures of DNA have been potentially mismodeled as pure Watson−Crick bps. Sites that experience significant Hoogsteen breathing can also be difficult to resolve by crystallography, as the elevated local dynamics could contribute to the lack of features in the local electron density49 or be suppressed by cryocooling of the crystals that is routinely performed to improve the resolution of the crystal structure.50 Conversely, there is also a need for methods that can independently verify Hoogsteen bps observed in crystal structures of DNA. For example, AT-rich duplexes composed entirely of Hoogsteen bps based on X-ray crystallography have been shown to form regular Watson−Crick bps when examined by solution NMR.15 Here, another potential example is given by the structure of the integration host factor (IHF) protein in complex with DNA13 (Figure 1B). IHF is a bacterial DNA binding protein that aids in the compaction of prokaryotic genomes by inducing significant DNA bending.51 It plays important roles in site-specific recombination of λ phage into the bacterial host genes,52 enhancing bacterial DNA replication,53 regulation of gene expression,54,55 and Cas1−Cas2-mediated spacer integration.56 IHF induces a global conformational change in the DNA duplex (i.e., ∼160° bending) upon binding, resulting in two sharp major groove-directed kinks separated by ∼7 Watson− Crick bps in pseudosymmetry-related sites (Figure 1B). In the crystal structure [Protein Data Bank (PDB) entry 1IHF], a nick between T48 and G49 (Figure 1B) was introduced at a kinked site to aid crystallization.13 Next to this site is an unusual Hoogsteen-like A21-T48 bp (Figure 1B). The adenine base adopts an anti rather than syn conformation that is uncharacteristic of Hoogsteen bps observed so far in duplexes.

Hoogsteen bps are also observed in some X-ray structures of DNA−protein complexes where they are thought to play roles in DNA shape recognition. 13,40−42 For example, two consecutive G-C+ Hoogsteen bps are observed in the complex between DNA and the TATA box binding protein.41 In one of these Hoogsteen bps, the syn guanine base appears to alleviate a steric clash between the guanine amino group and a nearby leucine side chain.41 In addition, one of the syn guanines partially stacks with the phenylalanine residue that inserts between the two G-C Hoogsteen bps.22 In the structure of DNA in complex with the DNA binding domain of p53 tumor suppressor protein, two consecutive A-T Hoogsteen bps in the consensus sequence contribute to a narrowed and more negatively charged minor groove in the flanking regions, which may favor electrostatic interactions with a positively charged arginine residue.42 Thus, proteins seem to employ a variety of mechanisms for interacting with Hoogsteen bps. Considering that it is now well established that Hoogsteen bps form robustly in naked duplex DNA23,35,40 with a preference for sites of major groove kinking,10,11 it comes as something of a surprise that Hoogsteen bps have not been more widely observed in structures of protein−DNA complexes, where structural distortions could destabilize Watson−Crick bps. Indeed, recent studies of DNA−drug complexes show that recognition of DNA can result in a variety of behaviors; some bps predominantly (>90% populations) adopt Watson−Crick or Hoogsteen conformations, while others having high (∼10%) fractional Hoogsteen populations relative to the naked DNA population (99.9% of the desired buffer. NMR Experiments. NMR data were collected on a 600 MHz Bruker NMR spectrometer equipped with an HCN cryogenic probe, a 600 MHz Bruker Avance III spectrometer equipped with an HCPN cryogenic probe, a 700 MHz Bruker Avance III spectrometer equipped with a triple-resonance HCN cryogenic probe, and an 800 MHz Varian DirectDrive2 spectrometer equipped with a triple-resonance HCN cryogenic probe. Data were processed and analyzed using NMR pipe57 and SPARKY (T. D. Goddard and D. G. Kneller, SPARKY 3, University of California, San Francisco), respectively. Chemical shift data were obtained using both TROSY and conventional two-dimensional (2D) HSQC experiments; resonance assignments were analyzed using 15N-edited 2D [1H, 1H] NOESY (mixing time of 180 ms), 2D H1′-C1′-AN9/TN1 HCN, and conventional HSQC experiments with broadband 15N decoupling and selective decoupling on purine-N9 or pyrimidine-N1. Spectra were recorded at 25 °C unless otherwise stated. The resonances in the IHF−DNA complex could be unambiguously assigned for single-site labeled samples (H′DNAA21 and H′-DNAA9). The aromatic resonances (A56-C8 and T48′-C6) in the IHF-bound H′-DNAA56T48 complex were assigned by turning on and off the selective carbon decoupling on T-C5 (∼110 ppm) in the conventional 2D HSQC experiment58 (data not shown). A56-C1′ and T48-C1′ in the free H′-DNAA56T48 were assigned by a 1H−15N 2D version of HCN experiment59 that correlates H1′ to adenine-N9 or thymine-N1 that has distinct chemical shifts (i.e., 170 ppm for adenine-N9 and 140 ppm for thymine-N1) (Figure S2A). However, the same HCN experiment could not detect signals in the DNA−protein complex due to significant transverse relaxation before acquisition. The A56-C1′ and T48-C1′ resonances in the DNA−protein complex were assigned by selectively decoupling adenine-N9 or thymine-N1, in comparison with the broadband 15N decoupling in a regular 2D HSQC experiment (Figure S2B). A weak NOE cross peak is observed that can be tentatively assigned to A21-H1′ in the 15N-edited NOESY spectra of the complex (see Figure 4B). Such an NOE is not expected for a regular Watson−Crick bp in which the distance between the protons is >6 Å, nor is it expected for the distorted Watson− Crick bp in the IHF protein crystal structure (>6 Å). Such a weak NOE could arise if T48 and A21 came into slightly closer proximity in solution, perhaps even transiently, when bound to IHF protein. This is consistent with observation of ∼0.22 ppm downfield shifted T48-H3 in the complex form relative to that in free DNA, which suggests stronger hydrogen bonding. It is also consistent with previously reported60 reduced imino proton exchange rates measured in a short 19 bp DNA upon binding to IHF. While a Hoogsteen bp could also bring these protons closer together (∼5.5 Å), we can rule out this possibility because we do not observe any evidence for the T48-H3−A21-H8 NOE (Figure S3). R1ρ RD experiments on the free H′-DNAA56T48 duplex were performed on the 700 MHz Bruker spectrometer as previously D

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Figure 2. NMR characterization of site-specifically 13C- and 15N-labeled H′-DNA duplexes. (A) H′-DNAA21, H′-DNAA56T48, and H′-DNAA9 sequences with 13C- and 15N-labeled nucleotides highlighted in color. (B) 1D 1H NMR spectra showing imino proton resonances in site-labeled DNA overlaid with those of unlabeled H′-DNA measured at 25 °C. (C) Overlay of 2D [13C, 1H] TROSY72 and 2D [15N, 1H] SOFAST HMQC73,74 NMR spectra measured for the three site-specifically labeled unbound H′-DNA sequences at 25 °C. Note that line broadening of A21C1′H1′ relative to other resonance is not observed in 2D HSQC (not TROSY) spectra (see Figure S3).

specific nucleotide positions were uniformly labeled with 13C and 15N (Figure 1B and Figure 2A). In one sample (H′DNAA21), the A21 that forms a Hoogsteen bp in the crystal structure was labeled (Figure 1B and Figure 2A). A second sample (H′-DNAA56T48) was prepared by labeling the A21 bp partner T48 as well as A56 at the pseudosymmetry-related site that forms a Watson−Crick bp in the crystal structure (Figure 1B and Figure 2A). A21, A56, and T48 are located at sites in which the DNA kinks sharply toward the major groove (Figure 1B and Figure S1). As a negative control, a third sample (H′DNAA9) was prepared labeling A9 that is located in a similar CAA sequence context but falls outside the consensus sequence in a region of the DNA structure that has a smaller degree of major groove kinking (Figure 1B and Figure S1). The buffer used for the NMR studies [25 mM HEPES, 100 mM NaCl, and 0.1 mM EDTA (pH 7.0)] was similar to crystallization buffer [10 mM HEPES, 100 mM NaCl, 0.1 mM EDTA, and 8% glycerol (pH 7.0)].68 We first analyzed the NMR spectra of the unbound DNA duplexes (Figure 2A). As expected, the one-dimensional (1D) 1 H spectra of all three DNA duplexes in the absence of IHF were essentially identical with well-resolved imino proton resonances that are consistent with a Watson−Crick B-form DNA duplex (Figure 2B). Excellent quality 2D NH and CH HSQC TROSY-based NMR spectra were obtained for all three duplexes clearly showing only a subset of resonances belonging to the labeled nucleotides (Figure 2C). The 1H, 13C, and 15N chemical shifts in the absence of IHF were all consistent with Watson−Crick pairing in B-form DNA. Resonances could readily be assigned in these site-labeled duplexes (see Methods and Figure S2). Assigning large nucleic acids (>20 kDa), such as the 34 bp DNA duplex, with conventional 2D/threedimensional NOESY experiments would be costly (∼1 mM for 13 C- and 15N-labeled samples) and challenging because of the low sensitivity and severe spectral overlap.69−71 In such cases, having a site-specifically labeled sample even at a low

anisotropy (A) was calculated on the basis of the parallel and perpendicular emission light intensities: A = (I − I⊥)/(I + 2I⊥)

where A is the measured anisotropy at a given concentration. The normalized anisotropy, Anorm, was obtained using A norm = (A − A 0)/(A max − A 0)

where A0 is the anisotropy in the absence of protein and Amax is the maximum measured anisotropy. Anorm was measured at varying protein concentrations (x) and then fitted to an equation describing single-site binding using the Levenberg− Marquardt nonlinear curve fitting algorithm implemented in the OriginPro 2016 software (OriginLab Corp.): ÄÅ ÉÑ Å Ñ A norm = B + CÅÅÅD + Kd + x − (D + Kd + x)2 − 4Dx ÑÑÑ ÅÇ ÑÖ where D is the constant fluorescein-labeled DNA concentration, B and C determine the anisotropies for free DNA and bound DNA, respectively, and Kd is the apparent dissociation constant. Because 7dA56 and 7dA21 are located in different strands, we kept the fluorescein-dT at the 5′-end of the opposing strand to the one containing the 7dA substitution (Figure 6A). Therefore, two control DNAs without 7-deaza substitution were made with the fluorescein label at either of the two strands (Figure 6). On the basis of the FP assay, the two control DNAs yielded similar Kd values (Kd1 = 1.0 ± 0.3 nM, and Kd2 = 1.4 ± 0.4 nM) for binding of unmodified H-DNA duplexes to IHF (Figure S4).

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RESULTS NMR Characterization of the Site-Specifically Labeled DNA Duplex. Three site-specifically labeled 34 bp DNA duplexes were prepared (see Methods) to study the IHF− DNA complex (Figure 1B and Figure 2A). In each case, E

DOI: 10.1021/acs.biochem.9b00027 Biochemistry XXXX, XXX, XXX−XXX

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Figure 3. Off-resonance relaxation dispersion profiles for unbound H′-DNAA56T48. Shown are off-resonance profiles as a function of the spin-lock offset (Ω 2π−1 kHz, where Ω = Ωobs − ωRF) and power (ωSL 2π−1 Hz, color-coded as indicated in the inset) measured at 37 °C. Error bars represent experimental uncertainties estimated from monoexponential fitting of n = 2 (A56-C1′) and n = 2 (T48-C1′) independently measured peak intensities using a Monte Carlo-based method (Methods). The solid line represents a fit to two-state exchange.35

Figure 4. NMR analysis of IHF−DNA complex formation. (A) 1D 1H spectra showing imino proton resonances in the IHF−H′-DNAA56T48 complex overlaid with those of unbound DNA. (B) 2D [13C, 1H] TROSY HSQC spectra of IHF−DNA complexes (colored) overlaid with those of unbound DNA (gray). Chemical shift perturbations are indicated by the black arrows. (C) Overlay of 2D [15N, 1H] SOFAST HMQC spectra of the IHF−H′-DNAA56T48 complex (green) and unbound H′-DNAA56T48 (gray) at 37 °C. The minor unbound DNA population is observed in the spectrum consistent with slow exchange. Complex-only spectra are shown in Figure S3. (D) 2D 15N-edited [1H, 1H] NOESY spectrum of the IHF−H′-DNAA56T48 complex showing T48H3-A21H2 NOE cross peaks consistent with Watson−Crick pairing at the A21-T48 bp. The T48-H3A21-H1′ NOE assignment (labeled with an asterisk) is tentative, and we cannot rule out the possibility that the resonance assigned to A21-H1′ corresponds to other neighboring sugar or amino protons.

tion (Rex) to the intrinsic transverse relaxation rate (R2,int) of NMR resonances during a relaxation period when the spin is irradiated with an applied radiofrequency (RF) pulse with varying powers (ωSL) and frequencies (ωRF). Experiments were performed at 37 °C. Indeed, we observed the expected RD at A56-C1′ (Figure 3). Fitting of the data to a two-state exchange model yields exchange parameters (pB = 0.3 ± 0.1%, kex = 1611 ± 525 s−1, and Δω = 4.2 ± 0.3 ppm) that are consistent with chemical exchange directed to Hoogsteen bps.35,38 The measured pB falls within the range (0.2−0.3%) measured in short (∼12 bp) DNA duplexes containing the same CAA trinucleotide sequence context at 30.5 °C.35 The measured exchange rate (kex = 1611 ± 525 s−1) is only slightly slower (∼3000−5000

concentration (∼0.3 mM in this study) provides a convenient methodology in a targeted approach to identify Watson− Crick/Hoogsteen bps in large nucleic acid molecules, potentially at lower external magnetic field strengths (i.e., 400 MHz) and noncryogenic probes. Transient Hoogsteen bps in Long Naked DNA Duplexes. Thus far, studies have examined transient Hoogsteen bps in relatively short (99%) at A21 but do not rule out having a small portion of the Hoogsteen population in equilibrium with the major Watson−Crick conformation. The small differences observed for the two sites (A21 and A56) are also consistent with small differences in NMR chemical shifts at these two bps in the IHF−DNA complex (Figure 4B).

Figure 5. Comparison of measured and calculated chemical shifts for Watson−Crick and Hoogsteen models in the IHF−DNA complex. Shown is the difference between chemical shifts [Δω = ωAix − ωA9x, in which i refers to different nucleotides (A21 or A56) and x to different types of nuclei (e.g., C1′, C8, and C2)]. Residue A9 is used as an internal reference. Shown are AFQM/MM-predicted values at A56 for 100% Hoogsteen (gray) and 100% Watson−Crick (orange) conformations and values measured by NMR for A21 (cyan) and A56 (green).

ensemble of conformations as done previously.39 Nevertheless, better agreement was observed between measured chemical shifts for both A21 and A56 and values calculated for the distorted Watson−Crick bp as in the X-ray structure as compared to the modeled Hoogsteen bp (Figure 5). While the agreement is somewhat better for the Hoogsteen form for AC8 and no distinctions can be made for AH1′ and AH2, AC1′, AC2, and AH8, resonances suggest better overall agreement with the Watson−Crick conformation. The chemical shift data therefore indicate that for the intact IHF−DNA complex in solution, the predominant conformation adopted by both A21T48 and A56-T13 is similar to that observed for the distorted A56-T13 Watson−Crick bp in the X-ray structure at the site without the nick. Indeed, this observed result is consistent with the X-ray structure of DNA in complex with Hbb, an IHFfamily bacterial DNA-bending protein. Although Hbb recognizes a different DNA sequence from IHF, the two protein−DNA complexes present similar overall highly bent DNA structures upon binding and two distorted A-T Watson−

Figure 6. Impact of 7-deazapurine substitutions on IHF−DNA binding affinity. (A) Sequences of DNA duplexes containing 7-deaza-dA modifications (red circles) and the fluorescein-dT label (green stars) at the 5′-end of the unmodified strand. Also shown is the chemical structure of 7-deaza-dA and its destabilization impact on Hoogsteen H-bonding. (B) Comparison of FP binding curves for 7-deaza-dA-modified DNA with its unmodified counterpart (control DNA). Control DNA refers to the unmodified DNA duplex with the fluorescein-dT label at the same position as the modified duplex. Data are fit to the single-site binding equation (see Methods). The error represents the standard deviation from triplicate (left) or duplicate (right) measurements. Fitted Kd and uncertainty values shown on the right. H

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DISCUSSION Resolving Hoogsteen versus Watson−Crick bps by X-ray crystallography is not always straightforward. Here, we demonstrated an NMR approach for examining Watson− Crick to Hoogsteen dynamics in large protein−DNA complexes. Application to the IHF−DNA complex indicates that A21-T48 adopts a predominantly Watson−Crick conformation and that the Hoogsteen bp observed in the crystal structure most likely arises due to the introduction of a nick near the kinked site. In this regard, it is interesting to note that Hoogsteen bps have been observed in structural contexts in which the nucleotide is not chemically linked to both neighbors, including bps at duplex termini, and also replication bps in the polymerase active site during replication.10,49 The NMR strategy presented here could facilitate studies of Hoogsteen bps in such contexts and possibly illuminate their biochemical consequences. In a recent study, it was shown that the binding of the echinomycin antibiotic to DNA increased the Hoogsteen population to 8%.22 Our data cannot rule out the possibility that the transient Hoogsteen population for A21-T48 in the IHF−DNA complex is also elevated by comparable amounts. While we observed line broadening for A21-C1′ and A56-C1′ relative to A9 and T48 in the IHF−DNA complex, further studies are needed to resolve whether these differences reflect microsecond-to-millisecond chemical exchange at A21-C1′ and A56-C1′ or faster picosecond-to-nanosecond dynamics that lead to narrowing of A9-C1′ and T48-C1′. TROSY-based NMR RD experiments combined with the site-specific labeling strategy used here may help extend the methodology to allow the study of transient Hoogsteen bps in large protein−DNA complexes.72,84−86 These studies could illuminate the DNA bending motions reported previously in the IHF−DNA complex87 as well as other dynamics that could impact the kinetic rates of DNA and IHF assembly.87,88 While our data suggest that the bps in the IHF−DNA complex form a predominantly Watson−Crick conformation, recent solution-state studies of the p53−DNA complex provide strong evidence for a predominantly Hoogsteen conformation that was observed in crystallographic structures.42,89 These crystal structures did not feature any nicks in the DNA. In particular, inosine or 2-oxoadenine substitutions that weaken or enhance Hoogsteen bp formation, as verified by X-ray crystallography, had more dramatic (∼6-fold) effects on DNA−protein binding affinities.89 This strongly suggests that Hoogsteen bps can form in protein−DNA complexes under solution conditions where they can contribute to shape recognition. The NMR strategy presented in this work opens the door for characterizing the dynamics of DNA in protein−DNA complexes. Indeed, very few studies60,90,91 have examined in atomic detail the dynamics of DNA when in complex with proteins and the potential role of these fluctuations in the stability and function of the complex. This approach can be applied to systems as large as 50 kDa but could potentially be extended to much larger complexes with the use of solid-state NMR. The site labeling approach also provides an important avenue for cataloguing valuable chemical shift−structure relationships that can be harnessed in the future to aid chemical shift-based characterization of DNA structure and dynamics.

Article

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.9b00027. Supplementary figures, supplementary table, and references (PDF) Accession Codes

P0A6X7 and P0A6Y1.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Huiqing Zhou: 0000-0002-9220-4742 David A. Case: 0000-0003-2314-2346 Present Address @

H.Z.: Institute for Biophysical Dynamics, The University of Chicago, Chicago, IL 60637. Author Contributions

H.M.A.-H. and H.Z. conceived the ideas for the project and designed experiments. H.Z. performed the NMR sample preparation, NMR, and biochemical experiments. B.S. helped set up the 15N-edited NOESY experiment for the protein− DNA complex. A.S. assisted in collecting FP data. Y. Xu helped collect R1ρ data for the long DNA duplex. Y. Xue assisted in molecular dynamics simulations of the IHF−DNA complex. Y. Z.P. and P.A.R. prepared the purified IHF protein. D.A.C. performed AFQM/MM calculations on free DNA and the DNA−protein complex. H.M.A.-H. and H.Z. wrote the manuscript with critical input from B.S., A.S., Y. Xu, Y. Xue, Y.Z.P., P.A.R., and D.A.C. Funding

This work was supported by National Institutes of Health Grant R01GM089846 to H.M.A.-H. Notes

The authors declare the following competing financial interest(s): H.M.A.-H. is an advisor to and holds an ownership interest in Nymirum Inc., an RNA-based drug discovery company.

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ACKNOWLEDGMENTS The authors thank Dr. Evgenia Nikolova, Dr. Janghyun Lee, Dr. Yu Chen, Dr. Isaac Kimsey, Honglue Shi, Atul Rangadurai, Dr. Mary Clay, Dr. Anisha Shakya, and Nicole Orlovsky for assistance and insightful discussions. The authors acknowledge technical support and resources from the Duke Magnetic Resonance Spectroscopy Center and the Duke Computer Cluster.

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REFERENCES

(1) Watson, J. D., and Crick, F. H. C. (1953) Molecular Structure of Nucleic Acids: A Structure for Deoxyribose Nucleic Acid. Nature 171, 737−738. (2) Hoogsteen, K. (1959) The structure of crystals containing a hydrogen-bonded complex of 1-methylthymine and 9-methyladenine. Acta Crystallogr. 12, 822−823. (3) Hoogsteen, K. (1963) The crystal and molecular structure of a hydrogen-bonded complex between 1-methylthymine and 9-methyladenine. Acta Crystallogr. 16, 907−916.

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duplex DNA using a cross-linked protein-DNA system. Nucleic Acids Res. 38, 4415−4425. (25) Freudenthal, B. D., Beard, W. A., Perera, L., Shock, D. D., Kim, T., Schlick, T., and Wilson, S. H. (2015) Uncovering the polymeraseinduced cytotoxicity of an oxidized nucleotide. Nature 517, 635−639. (26) Yang, H., and Lam, S. L. (2009) Effect of 1-methyladenine on thermodynamic stabilities of double-helical DNA structures. FEBS Lett. 583, 1548−1553. (27) Ling, H., Boudsocq, F., Plosky, B. S., Woodgate, R., and Yang, W. (2003) Replication of a cis-syn thymine dimer at atomic resolution. Nature 424, 1083−1087. (28) Nair, D. T., Johnson, R. E., Prakash, S., Prakash, L., and Aggarwal, A. K. (2004) Replication by human DNA polymerase-iota occurs by Hoogsteen base-pairing. Nature 430, 377−380. (29) Nair, D. T., Johnson, R. E., Prakash, L., Prakash, S., and Aggarwal, A. K. (2006) Hoogsteen base pair formation promotes synthesis opposite the 1,N6-ethenodeoxyadenosine lesion by human DNA polymerase [igr]. Nat. Struct. Mol. Biol. 13, 619−625. (30) Johnson, R. E., Prakash, L., and Prakash, S. (2005) Biochemical evidence for the requirement of Hoogsteen base pairing for replication by human DNA polymerase iota. Proc. Natl. Acad. Sci. U. S. A. 102, 10466−10471. (31) Makarova, A. V., and Kulbachinskiy, A. V. (2012) Structure of human DNA polymerase iota and the mechanism of DNA synthesis. Biochemistry (Moscow) 77, 547−561. (32) Palmer, A. G., III. (2014) Chemical exchange in biomacromolecules: past, present, and future. J. Magn. Reson. 241, 3−17. (33) Sekhar, A., and Kay, L. E. (2013) NMR paves the way for atomic level descriptions of sparsely populated, transiently formed biomolecular conformers. Proc. Natl. Acad. Sci. U. S. A. 110, 12867− 12874. (34) Xue, Y., Kellogg, D., Kimsey, I. J., Sathyamoorthy, B., Stein, Z. W., McBrairty, M., and Al-Hashimi, H. M. (2015) Chapter Two: Characterizing RNA Excited States Using NMR Relaxation Dispersion. In Methods in Enzymology (Sarah, A. W., and Frédéric, H. T. A., Eds.) pp 39−73, Academic Press. (35) Nikolova, E. N., Kim, E., Wise, A. A., O’Brien, P. J., Andricioaei, I., and Al-Hashimi, H. M. (2011) Transient Hoogsteen base pairs in canonical duplex DNA. Nature 470, 498−502. (36) Nikolova, E. N., Gottardo, F. L., and Al-Hashimi, H. M. (2012) Probing Transient Hoogsteen Hydrogen Bonds in Canonical Duplex DNA Using NMR Relaxation Dispersion and Single-Atom Substitution. J. Am. Chem. Soc. 134, 3667−3670. (37) Nikolova, E. N., Goh, G. B., Brooks, C. L., III, and Al-Hashimi, H. M. (2013) Characterizing the protonation state of cytosine in transient G•C Hoogsteen base pairs in duplex DNA. J. Am. Chem. Soc. 135, 6766−6769. (38) Alvey, H. S., Gottardo, F. L., Nikolova, E. N., and Al-Hashimi, H. M. (2014) Widespread transient Hoogsteen base pairs in canonical duplex DNA with variable energetics. Nat. Commun. 5, 4786. (39) Shi, H., Clay, M. C., Rangadurai, A., Sathyamoorthy, B., Case, D. A., and Al-Hashimi, H. M. (2018) Atomic structures of excited state A−T Hoogsteen base pairs in duplex DNA by combining NMR relaxation dispersion, mutagenesis, and chemical shift calculations. J. Biomol. NMR 70, 229−244. (40) Nikolova, E. N., Zhou, H., Gottardo, F. L., Alvey, H. S., Kimsey, I. J., and Al-Hashimi, H. M. (2013) A historical account of hoogsteen base-pairs in duplex DNA. Biopolymers 99, 955−968. (41) Patikoglou, G. A., Kim, J. L., Sun, L., Yang, S. H., Kodadek, T., and Burley, S. K. (1999) TATA element recognition by the TATA box-binding protein has been conserved throughout evolution. Genes Dev. 13, 3217−3230. (42) Kitayner, M., Rozenberg, H., Rohs, R., Suad, O., Rabinovich, D., Honig, B., and Shakked, Z. (2010) Diversity in DNA recognition by p53 revealed by crystal structures with Hoogsteen base pairs. Nat. Struct. Mol. Biol. 17, 423−429. (43) Pfoh, R., Cuesta-Seijo, J. A., and Sheldrick, G. M. (2009) Interaction of an echinomycin-DNA complex with manganese ions. Acta Crystallogr., Sect. F: Struct. Biol. Cryst. Commun. 65, 660−664.

(4) Pauling, L., and Corey, R. B. (1956) Specific hydrogen-bond formation between pyrimidines and purines in deoxyribonucleic acids. Arch. Biochem. Biophys. 65, 164−181. (5) Haschemeyer, A. E., and Sobell, H. M. (1963) The Crystal Structure of an Intermolecular Nucleoside Complex: Adenosine and 5-Bromouridine. Proc. Natl. Acad. Sci. U. S. A. 50, 872−877. (6) Sobell, H. M., Tomita, K. I., and Rich, A. (1963) The crystal structure of an intermolecular complex containing a guanine and a cytosine derivative. Proc. Natl. Acad. Sci. U. S. A. 49, 885−892. (7) Haschemeyer, A. E., and Sobell, H. M. (1965) The crystal structure of a hydrogen bonded complex of deoxyguanosine and 5bromodeoxycytidine. Acta Crystallogr. 19, 125−130. (8) Haschemeyer, A. E., and Sobell, H. M. (1965) The crystal structure of a hydrogen bonded complex of adenosine and 5bromouridine. Acta Crystallogr. 18, 525−532. (9) Wang, A. H., Ughetto, G., Quigley, G. J., Hakoshima, T., van der Marel, G. A., van Boom, J. H., and Rich, A. (1984) The molecular structure of a DNA-triostin A complex. Science 225, 1115−1121. (10) Zhou, H., Hintze, B. J., Kimsey, I. J., Sathyamoorthy, B., Yang, S., Richardson, J. S., and Al-Hashimi, H. M. (2015) New insights into Hoogsteen base pairs in DNA duplexes from a structure-based survey. Nucleic Acids Res. 43, 3420−3433. (11) Sathyamoorthy, B., Shi, H., Zhou, H., Xue, Y., Rangadurai, A., Merriman, D. K., and Al-Hashimi, H. M. (2017) Insights into Watson−Crick/Hoogsteen breathing dynamics and damage repair from the solution structure and dynamic ensemble of DNA duplexes containing m1A. Nucleic Acids Res. 45, 5586−5601. (12) Quigley, G. J., Ughetto, G., van der Marel, G. A., van Boom, J. H., Wang, A. H., and Rich, A. (1986) Non-Watson-Crick G.C and A.T base pairs in a DNA-antibiotic complex. Science 232, 1255−1258. (13) Rice, P. A., Yang, S., Mizuuchi, K., and Nash, H. A. (1996) Crystal structure of an IHF-DNA complex: a protein-induced DNA U-turn. Cell 87, 1295−1306. (14) Abrescia, N. G., Thompson, A., Huynh-Dinh, T., and Subirana, J. A. (2002) Crystal structure of an antiparallel DNA fragment with Hoogsteen base pairing. Proc. Natl. Acad. Sci. U. S. A. 99, 2806−2811. (15) Abrescia, N. G., González, C., Gouyette, C., and Subirana, J. A. (2004) X-ray and NMR studies of the DNA oligomer d(ATATAT): Hoogsteen base pairing in duplex DNA. Biochemistry 43, 4092−4100. (16) De Luchi, D., Tereshko, V., Gouyette, C., and Subirana, J. A. (2006) Structure of the DNA coiled coil formed by d(CGATATATATAT). ChemBioChem 7, 585−587. (17) Pous, J., Urpí, L., Subirana, J. A., Gouyette, C., Navaza, J., and Campos, J. L. (2008) Stabilization by extra-helical thymines of a DNA duplex with Hoogsteen base pairs. J. Am. Chem. Soc. 130, 6755−6760. (18) Singh, U. C., Pattabiraman, N., Langridge, R., and Kollman, P. A. (1986) Molecular mechanical studies of d(CGTACG)2: complex of triostin A with the middle A - T base pairs in either Hoogsteen or Watson-Crick pairing. Proc. Natl. Acad. Sci. U. S. A. 83, 6402−6406. (19) Ughetto, G., Wang, A. H. J., Quigley, G. J., van der Marel, G. A., van Boom, J. H., and Rich, A. (1985) A comparison of the structure of echinomycin and triostin A complexed to a DNA fragment. Nucleic Acids Res. 13, 2305−2323. (20) Gao, X., and Patel, D. J. (1988) NMR studies of echinomycin bisintercalation complexes with d(A1-C2-G3-T4) and d(T1-C2-G3A4) duplexes in aqueous solution: sequence-dependent formation of Hoogsteen A1.cntdot.T4 and Watson-Crick T1.cntdot.A4 base pairs flanking the bisintercalation site. Biochemistry 27, 1744−1751. (21) Cuesta-Seijo, J. A., and Sheldrick, G. M. (2005) Structures of complexes between echinomycin and duplex DNA. Acta Crystallogr., Sect. D: Biol. Crystallogr. 61, 442−448. (22) Xu, Y., McSally, J., Andricioaei, I., and Al-Hashimi, H. M. (2018) Modulation of Hoogsteen dynamics on DNA recognition. Nat. Commun. 9, 1473. (23) Yang, H., Zhan, Y., Fenn, D., Chi, L. M., and Lam, S. L. (2008) Effect of 1-methyladenine on double-helical DNA structures. FEBS Lett. 582, 1629−1633. (24) Lu, L., Yi, C., Jian, X., Zheng, G., and He, C. (2010) Structure determination of DNA methylation lesions N1-meA and N3-meC in J

DOI: 10.1021/acs.biochem.9b00027 Biochemistry XXXX, XXX, XXX−XXX

Article

Biochemistry (44) Wang, J. (2005) DNA polymerases: Hoogsteen base-pairing in DNA replication? Nature 437, E6−E7. (45) Aishima, J., Gitti, R. K., Noah, J. E., Gan, H. H., Schlick, T., and Wolberger, C. (2002) A Hoogsteen base pair embedded in undistorted B-DNA. Nucleic Acids Res. 30, 5244−5252. (46) Ethayathulla, A. S., Tse, P. W., Monti, P., Nguyen, S., Inga, A., Fronza, G., and Viadiu, H. (2012) Structure of p73 DNA-binding domain tetramer modulates p73 transactivation. Proc. Natl. Acad. Sci. U. S. A. 109, 6066−6071. (47) Kirouac, K. N., and Ling, H. (2011) Unique active site promotes error-free replication opposite an 8-oxo-guanine lesion by human DNA polymerase iota. Proc. Natl. Acad. Sci. U. S. A. 108, 3210−3215. (48) Nair, D. T., Johnson, R. E., Prakash, L., Prakash, S., and Aggarwal, A. K. (2005) Human DNA Polymerase ι Incorporates dCTP Opposite Template G via a G.C+ Hoogsteen Base Pair. Structure 13, 1569−1577. (49) Hintze, B. J., Richardson, J. S., and Richardson, D. C. (2017) Mismodeled purines: implicit alternates and hidden Hoogsteens. Acta Crystallographica Section D 73, 852−859. (50) Fenwick, R. B., van den Bedem, H., Fraser, J. S., and Wright, P. E. (2014) Integrated description of protein dynamics from roomtemperature X-ray crystallography and NMR. Proc. Natl. Acad. Sci. U. S. A. 111, E445. (51) Swinger, K. K., and Rice, P. A. (2004) IHF and HU: flexible architects of bent DNA. Curr. Opin. Struct. Biol. 14, 28−35. (52) Weisberg, R. A., Enquist, L. W., Foeller, C., and Landy, A. (1983) Role for DNA homology in site-specific recombination: The isolation and characterization of a site affinity mutant of coliphage λ. J. Mol. Biol. 170, 319−342. (53) Gamas, P., Burger, A. C., Churchward, G., Caro, L., Galas, D., and Chandler, M. (1986) Replication of pSC101: effects of mutations in the E. coli DNA binding protein IHF. Molecular and General Genetics MGG 204, 85−89. (54) Goosen, N., van Heuvel, M., Moolenaar, G. F., and van de Putte, P. (1984) Regulation of Mu transposition II. The Escherichia coli HimD protein positively controls two repressor promoters and the early promoter of bacteriophage Mu. Gene 32, 419−426. (55) Friedman, D. I. (1988) Integration host factor: A protein for all reasons. Cell 55, 545−554. (56) Nuñez, J. K., Bai, L., Harrington, L. B., Hinder, T. L., and Doudna, J. A. (2016) CRISPR Immunological Memory Requires a Host Factor for Specificity. Mol. Cell 62, 824−833. (57) Delaglio, F., Grzesiek, S., Vuister, G. W., Zhu, G., Pfeifer, J., and Bax, A. (1995) NMRPipe: A multidimensional spectral processing system based on UNIX pipes. J. Biomol. NMR 6, 277−293. (58) Ulrich, E. L., Akutsu, H., Doreleijers, J. F., Harano, Y., Ioannidis, Y. E., Lin, J., Livny, M., Mading, S., Maziuk, D., Miller, Z., Nakatani, E., Schulte, C. F., Tolmie, D. E., Kent Wenger, R., Yao, H., and Markley, J. L. (2007) BioMagResBank. Nucleic Acids Res. 36, D402−D408. (59) Sklenár,̌ V., Peterson, R. D., Rejante, M. R., and Feigon, J. (1993) Two-and three-dimensional HCN experiments for correlating base and sugar resonances in 15N, 13C-labeled RNA oligonucleotides. J. Biomol. NMR 3, 721−727. (60) Dhavan, G. M., Lapham, J., Yang, S., and Crothers, D. M. (1999) Decreased imino proton exchange and base-pair opening in the IHF-DNA complex measured by NMR. J. Mol. Biol. 288, 659− 671. (61) Hansen, A. L., Nikolova, E. N., Casiano-Negroni, A., and AlHashimi, H. M. (2009) Extending the range of microsecond-tomillisecond chemical exchange detected in labeled and unlabeled nucleic acids by selective carbon R1ρ NMR Spectroscopy. J. Am. Chem. Soc. 131, 3818−3819. (62) Kimsey, I. J., Petzold, K., Sathyamoorthy, B., Stein, Z. W., and Al-Hashimi, H. M. (2015) Visualizing transient Watson-Crick-like mispairs in DNA and RNA duplexes. Nature 519, 315−320.

(63) Swails, J., Zhu, T., He, X., and Case, D. A. (2015) AFNMR: automated fragmentation quantum mechanical calculation of NMR chemical shifts for biomolecules. J. Biomol. NMR 63, 125−139. (64) Richardson, W. H., Peng, C., Bashford, D., Noodleman, L., and Case, D. A. (1997) Incorporating solvation effects into density functional theory: Calculation of absolute acidities. Int. J. Quantum Chem. 61, 207−217. (65) Koster, A. M., Geudtner, G., Calaminici, P., Casida, M. E., Dominguez, V. D., Flores-Moreno, R., Gamboa, G. U., Goursot, A. H. T., Ipatov, A. J. F., del Campo, J. M., Reveles, J. U., Vela, A., ZunigaGutierrez, B., and Salahub, D. R. (2011) deMon2k, version 4, Cinvestav, Mexico City. (66) Handy, N. C., and Cohen, A. J. (2001) Left-right correlation energy. Mol. Phys. 99, 403−412. (67) Case, D. A. (2013) Chemical shifts in biomolecules. Curr. Opin. Struct. Biol. 23, 172−176. (68) Lynch, T. W., Read, E. K., Mattis, A. N., Gardner, J. F., and Rice, P. A. (2003) Integration Host Factor: Putting a Twist on Protein−DNA Recognition. J. Mol. Biol. 330, 493−502. (69) Lukavsky, P. J., Kim, I., Otto, G. A., and Puglisi, J. D. (2003) Structure of HCV IRES domain II determined by NMR. Nat. Struct. Mol. Biol. 10, 1033. (70) Lu, K., Heng, X., Garyu, L., Monti, S., Garcia, E. L., Kharytonchyk, S., Dorjsuren, B., Kulandaivel, G., Jones, S., Hiremath, A., Divakaruni, S. S., LaCotti, C., Barton, S., Tummillo, D., Hosic, A., Edme, K., Albrecht, S., Telesnitsky, A., and Summers, M. F. (2011) NMR Detection of Structures in the HIV-1 5′-Leader RNA That Regulate Genome Packaging. Science 334, 242. (71) Dieckmann, T., and Feigon, J. (1997) Assignment methodology for larger RNA oligonucleotides: Application to an ATP-binding RNA aptamer. J. Biomol. NMR 9, 259−272. (72) Pervushin, K., Riek, R., Wider, G., and Wüthrich, K. (1997) Attenuated T2 relaxation by mutual cancellation of dipole−dipole coupling and chemical shift anisotropy indicates an avenue to NMR structures of very large biological macromolecules in solution. Proc. Natl. Acad. Sci. U. S. A. 94, 12366−12371. (73) Farjon, J., Boisbouvier, J., Schanda, P., Pardi, A., Simorre, J.-P., and Brutscher, B. (2009) Longitudinal-relaxation-enhanced NMR experiments for the study of nucleic acids in solution. J. Am. Chem. Soc. 131, 8571−8577. (74) Sathyamoorthy, B., Lee, J., Kimsey, I., Ganser, L. R., and AlHashimi, H. (2014) Development and application of aromatic [13C, 1H] SOFAST-HMQC NMR experiment for nucleic acids. J. Biomol. NMR 60, 77−83. (75) Palmer, A. G., III (2016) A dynamic look backward and forward. J. Magn. Reson. 266, 73−80. (76) Yang, C.-C., and Nash, H. A. (1989) The interaction of E. coli IHF protein with its specific binding sites. Cell 57, 869−880. (77) Ozers, M. S., Hill, J. J., Ervin, K., Wood, J. R., Nardulli, A. M., Royer, C. A., and Gorski, J. (1997) Equilibrium Binding of Estrogen Receptor with DNA Using Fluorescence Anisotropy. J. Biol. Chem. 272, 30405−30411. (78) Yang, S. W., and Nash, H. A. (1995) Comparison of protein binding to DNA in vivo and in vitro: defining an effective intracellular target. EMBO J. 14, 6292−6300. (79) Wang, S., Cosstick, R., Gardner, J. F., and Gumport, R. I. (1995) The specific binding of Escherichia coli integration host factor involves both major and minor grooves of DNA. Biochemistry 34, 13082−13090. (80) Gilbert, D. E., van der Marel, G. A., van Boom, J. H., and Feigon, J. (1989) Unstable Hoogsteen base pairs adjacent to echinomycin binding sites within a DNA duplex. Proc. Natl. Acad. Sci. U. S. A. 86, 3006−3010. (81) Mouw, K. W., and Rice, P. A. (2007) Shaping the Borrelia burgdorferi genome: crystal structure and binding properties of the DNA-bending protein Hbb. Mol. Microbiol. 63, 1319−1330. (82) Ganguly, M., Wang, F., Kaushik, M., Stone, M. P., Marky, L. A., and Gold, B. (2007) A study of 7-deaza-2′-deoxyguanosine−2′deoxycytidine base pairing in DNA. Nucleic Acids Res. 35, 6181−6195. K

DOI: 10.1021/acs.biochem.9b00027 Biochemistry XXXX, XXX, XXX−XXX

Article

Biochemistry (83) Kowal, E. A., Ganguly, M., Pallan, P. S., Marky, L. A., Gold, B., Egli, M., and Stone, M. P. (2011) Altering the Electrostatic Potential in the Major Groove: Thermodynamic and Structural Characterization of 7-Deaza-2′-deoxyadenosine:dT Base Pairing in DNA. J. Phys. Chem. B 115, 13925−13934. (84) Korzhnev, D. M., Kloiber, K., Kanelis, V., Tugarinov, V., and Kay, L. E. (2004) Probing Slow Dynamics in High Molecular Weight Proteins by Methyl-TROSY NMR Spectroscopy: Application to a 723-Residue Enzyme. J. Am. Chem. Soc. 126, 3964−3973. (85) Igumenova, T. I., and Palmer, A. G. (2006) Off-Resonance TROSY-Selected R1ρ Experiment with Improved Sensitivity for Medium- and High-Molecular-Weight Proteins. J. Am. Chem. Soc. 128, 8110−8111. (86) Kitevski-LeBlanc, J. L., Yuwen, T., Dyer, P. N., Rudolph, J., Luger, K., and Kay, L. E. (2018) Investigating the Dynamics of Destabilized Nucleosomes Using Methyl-TROSY NMR. J. Am. Chem. Soc. 140, 4774−4777. (87) Vivas, P., Velmurugu, Y., Kuznetsov, S. V., Rice, P. A., and Ansari, A. (2012) Mapping the Transition State for DNA Bending by IHF. J. Mol. Biol. 418, 300−315. (88) Sugimura, S., and Crothers, D. M. (2006) Stepwise binding and bending of DNA by Escherichia coli integration host factor. Proc. Natl. Acad. Sci. U. S. A. 103, 18510. (89) Golovenko, D., Bräuning, B., Vyas, P., Haran, T. E., Rozenberg, H., and Shakked, Z. (2018) New Insights into the Role of DNA Shape on Its Recognition by p53 Proteins. Structure 26, 1237−1250. (90) Fernandez, C., Szyperski, T., Ono, A., Iwai, H., Tate, S., Kainosho, M., and Wuthrich, K. (1998) NMR with 13C, 15N-doublylabeled DNA: the Antennapedia homeodomain complex with a 14mer DNA duplex. J. Biomol. NMR 12, 25−37. (91) Dupureur, C. M. (2006) Unique 31P Spectral Response to the Formation of a Specific Restriction Enzyme−DNA Complex. Nucleosides, Nucleotides Nucleic Acids 25, 747−764.

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