Molecular Mechanism, Dynamics, and Energetics of Protein-Mediated

Subscriber access provided by UNIV OF DURHAM

Article

Molecular Mechanism, Dynamics, and Energetics of Protein-mediated Dinucleotide Flipping in a Mismatched DNA: A Computational Study of RAD4-DNA Complex Kartheek Pitta, and Marimuthu Krishnan J. Chem. Inf. Model., Just Accepted Manuscript • DOI: 10.1021/acs.jcim.7b00636 • Publication Date (Web): 23 Feb 2018 Downloaded from http://pubs.acs.org on February 23, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Journal of Chemical Information and Modeling is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of Chemical Information and Modeling

Molecular Mechanism, Dynamics, and Energetics of Protein-mediated Dinucleotide Flipping in a Mismatched DNA: A Computational Study of RAD4-DNA complex Kartheek Pitta and Marimuthu Krishnan∗ Center for Computational Natural Sciences and Bioinformatics, International Institute of Information Technology, Gachibowli 500032, Hyderabad, Telangana, India E-mail: [email protected] Abstract DNA damages alter genetic information and adversely affect gene expression pathways leading to various complex genetic disorders and cancers. DNA repair proteins recognize and rectify DNA damages and mismatches with high fidelity. A critical molecular event that occurs during most protein-mediated DNA repair processes is the extrusion of orphaned bases at the damaged site facilitated by specific repairing enzymes. The molecular-level understanding of the mechanism, dynamics, and energetics of base extrusion is necessary to elucidate the molecular basis of protein-mediated DNA damage repair. The present article investigates the molecular mechanism of dinucleotide extrusion in a mismatched DNA (containing a stretch of three contiguous thymidine-thymidine base pairs) facilitated by Radiation sensitive 4 (RAD4), a key DNA repair protein, on an atom-by-atom basis using molecular dynamics (MD) and umbrella-sampling (US) simulations. Using atomistic models of RAD4-free and

1

ACS Paragon Plus Environment

Journal of Chemical Information and Modeling 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

RAD4-bound mismatched DNA, the free energy profiles associated with extrusion of mismatched partner bases are determined for both systems. The mismatched bases adopted the most stable intra-helical conformation and their extrusion was unfavorable in RAD4-free mismatched DNA due to the presence of prohibitively high barriers (> 12.0 kcal/mol) along the extrusion pathways. Upon binding of RAD4 to the DNA, the global free energy minimum is shifted to the extra-helical state indicating the key role of RAD4-DNA interactions in catalyzing the dinucleotide base extrusion in the DNA-RAD4 complex. The critical residues of RAD4 contributing to the conformational stability of the mismatched bases are identified and the energetics of insertion of a β-hairpin of RAD4 into the DNA duplex is examined. The conformational landscapebased mechanistic insight into RAD4-mediated base extrusion provided here may serve as a useful baseline to understand the molecular basis of xeroderma pigmentosum C (XPC)-mediated DNA damage repair in humans.

Introduction The integrity of genome is constantly challenged by DNA damages caused by various damaging agents including free radicals, UV radiation, oxidation, alkylation, genotoxic chemicals, viruses, and anti-cancer drugs . 1–3 To safeguard genetic information, biological cells have sophisticated survival mechanisms to recognize and rectify DNA damages with high fidelity . 4–11 The task of recognition and repair of damaged DNA is carried out by a group of specific proteins. 12,13 It is essential to understand the molecular basis of protein-mediated DNA damage repair to comprehend on the initiation and evolution of various diseases including a variety of complex genetic disorders and cancers. 14–20 In order to understand how specific proteins recognize and repair DNA damages, it is necessary to identify proteins that interact directly with the damaged DNA and investigate the nature of their interactions with DNA. 21,22 In this regard, X-ray diffraction (XRD) and solution NMR experiments 23 probe the crystal and solution structures, respectively, of

2


Page 2 of 40

Page 3 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


damaged DNA-protein complexes and offer the initial structural clues into the nature of DNA-protein interactions on an atom-by-atom basis.

Radiation sensitive 4 (RAD4) is a key protein that recognizes a variety of DNA damages in yeast cells. 24–30 As an orthologue of xeroderma pigmentosum C (XPC), which is a human DNA repair protein, with functional similarity, RAD4 serves as a model system to understand XPC-mediated DNA damage repair in humans. 31–34 Like XPC, RAD4 also coordinates with a group of downstream factors to recognize and remove damaged lesions and distortions in DNA. 35,36 Figure 1 illustrates crucial steps involved in RAD4-facilitated repair of DNA damages. 18,28,37–50 RAD4 initially scans the DNA and recognizes the undamaged partner nucleotides opposite to the lesion 41,42 (Figure 1(A)). Once the damage site is identified, RAD4 stalls and recruits specific proteins including transcription initiation factor IIH (TFIIH), xeroderma pigmentosum B and D (XPB and XPD) helicases to unwind both sides of the duplex DNA leading to a bubble formation consisting of about 20-30 nucleotides at the damage site 18,43,44 (Figure 1(B-D)). The two single-stranded DNA binding proteins, replication protein A1 (RPA) and xeroderma pigmentosum complementation group A (XPA) bind to and stabilize the single-stranded DNA 45 formed by the above mentioned helicase activity (Figure 1(E)). Subsequently, a 24-32-mer oligonucleotide in the damaged strand are cleaved on either side of the lesion by the endonucleases encoded by XPF and XPG genes 46–48 (Figure 1(F)). Finally, the single-stranded undamaged strand is synthesized and ligated by ligase-III and X-ray repair cross-complementing protein 1 (XRCC1) 49–52 (Figure 1(G)(H)). Although biochemical studies continue to advance our understanding of RAD4-DNA complexes, the atomic-level details of how RAD4 recognizes and repairs DNA damages and of the critical molecular interactions that govern the energetics of the repair process remain unclear. 28,42 XRD studies on the crystal structure of RAD4 bound to DNA containing a cyclobutane pyrimidine dimer (CPD) lesion superimposed on a stretch of three contiguous thymidine-thymidine mis-

3



Figure 1: A schematic illustration of the critical steps involved in RAD4-facilitated nucleotide excision repair mechanism of damaged DNA. (A) Damage recognition by RAD4. (B) BHD2 and BHD3 domains of RAD4 interact with the damaged DNA. (C) RAD4 extrudes the damaged bases. (D) Helicase activity by XPB, XPD and TFIIH. (E) XPA and RPA stabilize the ssDNA. (F) Endonuclease activity by XPF and XPG. (G) Polymerization by PCNA and ligation of DNA. (H) Repaired DNA. matches (hereafter, 3T mismatch) offer a structure-based mechanism of DNA damage repair by RAD4. 42 RAD4 contains three functionally-relevant β-hairpin domains (BHD): BHD1, BHD2 and BHD3 (shown in Figure 2). These domains play an important role in damage sensing and extrusion of mismatched thymine partners in RAD4-DNA complex. 53,54 BHD2 and BHD3 bind to the damaged part of the DNA, whereas BHD1 binds to the undamaged part of the DNA. The crystal structure of RAD4-DNA complex reveals that BHD3 inserts a hairpin loop into the lesion region and expels the two undamaged nucleotides opposite to the lesion out of the duplex, resulting in dinucleotide flipping from the intra-helical to

4


Page 4 of 40

Page 5 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


extra-helical conformation. 42 The expelled nucleotide bases are caught safely in a glove of a few specific residues of BHD2 and BHD3 of RAD4. This protein-induced flipping out of the mismatched bases at the site of damage is believed to be a key step in RAD4-mediated nucleotide excision repair (NER). Computational methods based on molecular dynamics (MD) simulation and enhanced sampling free energy methods have emerged as powerful tools to understand the energetics, dynamics, structural perturbations and molecular mechanisms associated with DNA repair processes. 55–57 The base flipping in DNA-protein complexes is thought to be an active process involving relatively high energy barriers and thus occurs on a time scale of milliseconds or longer. 58–60 These rare events are unlikely to occur in 10-100 nanosecond direct MD simulations. However, many independent trajectories obtained from multiple MD simulations can provide meaningful statistics on such rare events, but it is a computationally expensive exercise. 61,62 The accelerated MD simulations using external biasing potentials to enhance the conformational sampling can provide valuable insights into the energetics and mechanisms of DNA base flipping events. 63,64 The pseudo-dihedral angles, which describe the rotational dynamics of the chosen bases, are often used as reaction coordinates to determine the relative free energies of a spectrum of stable and metastable conformational states that lie between the intra- and extra-helical states of the chosen bases. 56,65 Moreover, the free energy profiles associated with DNA base flipping obtained from such enhanced sampling methods provide detailed insights into relative stabilities of different conformational states and the activation energy barriers for transitions between them.

Based on earlier studies on various protein-DNA complexes, the mechanism of base flipping during protein-mediated DNA damage repair processes is broadly classified into two categories: the (i) active and (ii) passive mechanisms. 53,66–69 The spontaneously flipped out bases are recognized and captured by the protein in the passive or conformational capture mechanism while the base flipping is initiated and controlled by the protein in the active or

5



correlated motion mechanism. 35,53,66–69 A recent computational study examined the binding free energy profile of RAD4 as it binds to a CPD-containing DNA duplex with mismatched thymine partner bases. They estimated the free energy barrier for the formation of the productively bound complex that matched the experimental crystal structure of Min and co-workers. The calculated free energy barrier of ∼10 kcal/mol for the conformational capture mechanism was observed to be higher than a barrier of ∼4 kcal/mol obtained for the correlated motion mechanism. The observed difference in free energy barriers indicated that the correlated motion mechanism, in which uT15 and uT16 flip in concert, is the preferred mechanism underpinning RAD4mediated DNA damage recognition for this CPD lesion in a mismatch context. 53

In the present article, we have used molecular dynamics and umbrella sampling simulations to determine the free energy surfaces associated with the RAD4-facilitated dinucleotide base flipping and to examine the relative stability of the RAD4-bound mismatched DNA with respect to the RAD4-free DNA. A detailed comparison of the free energy profiles of the RAD4-free and RAD4-bound mismatched DNA reveals that RAD4 binding-induced structural distortions in DNA near the site of mismatch result in a conformational shift of mismatched bases from their intra-helical to extra-helical states. The catalytic role of RAD4 in stabilizing the extra-helical conformations of mismatched partner bases underscores the important role of RAD4-DNA interactions in the dinucleotide base flipping.

Methods Model Systems The X-ray crystal structure of RAD4-DNA complex with 3T mismatch (PDB ID: 2QSH) was used as the starting structure for MD simulation of the complex. 42 A missing loop G(518)R(519)P(520)K(521)G(522)E(523)A(524)E(525) in the BHD2 domain of RAD4 and 6


Page 6 of 40

Page 7 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


the two missing disordered thymine bases in the damaged strand of DNA were modeled using the Modeller 70 and the PSFGEN tool of VMD, 71 respectively. The newly added residues and bases were initially subjected to energy minimization (5000 steps of steepest decent minimization followed by 5000 steps of conjugate gradient minimization) with all other atoms restrained to their crystallographic positions.

Simulation Details MD simulations were performed using the GROMACS 4.5.5 simulation package 72,73 with amber99bsc1 74 force field for protein and DNA and TIP3P force field for water molecules. The equations of motion were integrated using the velocity Verlet algorithm with a time step of 2 fs. The bond lengths involving hydrogen atoms were constrained using LINCS. 75 The periodic boundary conditions were employed along all directions. The particle mesh Ewald (PME) approach 76,77 with a direct space cut-off of 10 ˚ A was used for the long-range electrostatic interactions. The pressure control was achieved using the Parrinello-Rahman method 78,79 with a time constant of 1 ps and the temperature was controlled using a velocityrescale thermostat 80 with a time constant of 0.1 ps. The energy minimization for all the systems was done using the steepest descent (5000 steps) and conjugate gradient (5000 steps) methods. The equilibration of the energy minimized structures was carried out in three different stages: the initial stage involves NVT annealing from 0 K to 298 K for 300 ps with position restraints (with a harmonic spring constant of 239 kcal/mol/˚ A2 ) on DNA heavy atoms followed by NPT equilibration at 298 K and at 1 atm pressure for 1 ns. In the last stage of equilibration, all the position restraints were removed and each system was subjected to 1 ns equilibration followed by 20 ns NPT production run.

7



Umbrella Sampling The umbrella sampling (US) 81 method was used to quantify the base-flipping energetics in RAD4-free and RAD4-bound DNA. In particular, the present study examines the flipping energetics and dynamics of a pair of mismatched partner thymine bases (denoted by uT15 and uT16 (Figure 2(d))- here ’u’ stands for the undamaged strand of DNA) in the undamaged strand of DNA. uT15 and uT16 are also referred to as 50 and 30 partner thymine bases, respectively. 53 The pseudo-dihedral angles (Φ and Ψ shown in Figure 3(C)) involving the centers of mass of heavy atoms in the paired bases of interest (uT15 and uT16) and the connected ribose sugars were chosen as reaction coordinates or collective variables (CV). 56 Here, the reaction coordinates Ψ and Φ describe flipping dynamics of uT15 and uT16, respectively. The reaction coordinate Ψ was chosen such that a clockwise rotation (Ψ: 0o → 90o → 180o ) of uT15 about the pseudo-central bond between adjacent sugar moieties (refer Figure 3(C)) defines the major-groove extrusion pathway for uT15. For uT16, a counter-clockwise rotation (Φ: 0o → −90o → −180o ) defines its major-groove pathway in RAD4-DNA complex. Independent US calculations were performed for the individual reaction coordinates (Φ and Ψ) using the PLUMED2.2 82 plugin of GROMACS. The allowed range (-180o ≤ Φ ≤ 180o and -180o ≤ Ψ ≤ 180o ) of a given reaction coordinate was divided into 72 windows of width 5o and independent 3 ns NPT simulation with a harmonic biasing potential (with a force constant of 239 kcal/mol/rad2 ) was carried out for each window. The potentials of mean force (PMFs) (F(Φ) and F(Ψ)) were determined from the respective umbrella sampling trajectories using the weighted histogram analysis method (WHAM). 83 F(Φ) and F(Ψ) thus obtained provide detailed information on favorable conformations of the mismatched partner bases, their relative stabilities and the barriers between them along the chosen reaction coordinates. The convergence of the free energy profiles was examined by monitoring the time evolution

8


Page 8 of 40

Page 9 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


of a convergence quantity, η, which is defined as follows Φ=π 1 X η= [F (Φ, t) − F (Φ, t + δt)]2 N Φ=−π

(1)

where N is the number of windows used in the umbrella sampling, F(Φ, t) and F(Φ, t+∆ t) are the free energy profiles calculated at two adjacent time windows, t and t+∆t, respectively. The sampling time was chosen such that η decreases to a value close to zero to ensure a reasonable convergence of the free energy profiles. 84,85

Results and Discussion Free Energy Profiles The crystal structure of a CPD-containing RAD4-DNA complex reveals two important structural changes in DNA upon binding to RAD4; 42 firstly, the double helical structure of DNA is distorted to a kinked conformation with a bend at the lesion site (Figure 2(b)); secondly, the mismatched partner thymine bases (uT15 and uT16) opposite to the lesion are flipped out of the DNA duplex into the binding pocket of RAD4. In this section, we compare the free energy profiles associated with flipping of these mismatched thymine bases in RAD4-free and RAD4-bound mismatched DNA to understand the role of DNA-RAD4 interactions in the dinucleotide base flipping energetics and dynamics.

Base Flipping Energetics in the Absence of RAD4 Figure 4 shows the free energy profiles F(Φ) and F(Ψ) associated with extrusion of uT15 and uT16 obtained from the umbrella sampling simulations for BDI (i.e., RAD4-free DNA) and REC (i.e., RAD4-bound DNA) model systems. Hereafter, FBDI (Φ or Ψ) and FREC (Φ or Ψ) denote the calculated free energy profiles for the BDI and REC model systems, respectively. The time evolution of the reaction coordinates Φ and Ψ obtained from direct MD simulations

9



Figure 2: Two model systems of interest: (a) a double-helical B-DNA (BDI), (b) The conformation of DNA in the RAD4-DNA complex. (c) The crystal structure of RAD4-DNA complex (REC) with trans glutaminase domain (TGD) (orange), β-hairpin domains BHD1 (violet), BHD2 (blue), BHD3 (red) of RAD4 and the extruded mismatched (uT15 and uT16) bases (cyan) of DNA. ; (d) The sequence of DNA in these models with a 3T mismatch lesion. These images were made using pymol. is also shown in Figure 4 (A) and Figure 4 (B), respectively. In the absence of RAD4, FBDI (Φ) exhibits a global free energy minimum at Φ ∼14o

10


Page 10 of 40

Page 11 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 3: The definitions of (A) intra- and extra-helical conformations (B) the major- and minor-groove pathways for dinucleotide base extrusion (C) the pseudo-dihedral angles Φ and Ψ. The centers of mass of all the heavy atoms in the base pair (cyan ellipse), ribose sugars (green circles), and the dangling base to be flipped (red circle) define the pseudo-dihedral angles. surrounded by a steep energy well indicating that the intra-helical conformation is the most stable conformation for uT16 in BDI. FBDI (Φ) also reveals that the extra-helical conformations (with |Φ| >100o ) of uT16 are at least ∼10 kcal/mol higher in energy relative to the most-stable intra-helical conformation and that there are no noticeable free energy minima in the extra-helical region (| Φ |>100o ) of the conformational space of uT16. The higher relative stability of the intra-helical conformation of uT16 in BDI can be attributed to its inter-strand base-pairing hydrogen bonds and the stacking interactions with neighboring bases in the Watson-Crick helical geometry of the DNA duplex. The calculated activation barrier of ∼12.5 kcal/mol for the extrusion of uT16 via the major- and minor-groove pathways suggests that uT16 is unlikely to flip out in the absence of RAD4 at room temperature. FBDI (Ψ) exhibits a global energy minimum at Ψ ∼ 37o (Figure 4B) indicating that uT15 also prefers an intra-helical conformation in the absence of RAD4. The extra-helical conformations (| Ψ |>100o ) of uT15 are at least 5.5 kcal/mol (along the major-groove path) and 15 kcal/mol (along the minor-groove path) higher in energy relative to the most stable 11



20

A

14

15

10 8

10

6 4

Time (ns)

F(Φ) kcal/mol

12

5

2 0 -180 -150 -120 -90 -60 -30

0

0

30 60 90 120 150 180

Φ (degree) 18 16

20

B

14

15

12 10 10 8

Time (ns)

F(Ψ) kcal/mol

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 40

6 5

4 2 0 -180 -150 -120 -90 -60 -30

0

30

60

90

0 120 150 180

Ψ (degree) Figure 4: The umbrella sampling-derived PMF profiles associated with dinucleotide base extrusion in BDI (black) and REC (red) are shown along with the corresponding MD-derived time evolution of Φ and Ψ (BDI: green; REC: purple). intra-helical conformation at Ψ ∼ 37o . Here again, the absence of noticeable energy minima in the extra-helical conformational region and higher energies of the extra-helical conformations indicate that the extra-helical conformations are unfavorable for uT15 in BDI. The snapshots obtained from the umbrella sampling trajectories illustrating extrusion of uT15 and uT16 12


Page 13 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


via the major-groove and minor-groove pathways in RAD4-free mismatched DNA are shown in Figure S1 and Figure S2 (Supporting Information). A comparison of FBDI (Φ) and FBDI (Ψ) reveals that the free energy well around the global minimum in FBDI (Ψ) is relatively broader and more asymmetric than the steep energy well around the global minimum in FBDI (Φ). In particular, both FBDI (Φ) and FBDI (Ψ) are relatively shallower along their respective major groove pathways (Figure 4), but the degree of shallowness is more in FBDI (Ψ) than that in FBDI (Φ). The higher well width and higher degree of shallowness along the major groove pathway in FBDI (Ψ) suggests that uT15 is likely to be vibrationally more flexible in its intra-helical conformational state than that of uT16. The higher degree of shallowness along the major groove pathways relative to that for the minor groove pathways also suggests that both uT15 and uT16 are likely to prefer their respective major groove pathways for their extrusion in BDI (see Figure S5 and Figure S6 - Supporting Information). The calculated activation barriers along the major- and minor-groove pathways in BDI are ∼12.5 kcal/mol and ∼16 kcal/mol, respectively, for uT15 and ∼12.5 kcal/mol for uT16. Since the extrusion barriers along the major groove pathways of uT15 and uT16 are almost the same, it is unclear which of these bases flips first during the course of extrusion in BDI. However, a relatively higher barrier observed along the minor-groove pathway of uT15 suggests that, given sufficient thermal energy, uT16 is likely to extrude first followed by the extrusion of uT15 in BDI. Using a different pseudo-dihedral angle for the correlated flipping of both uT15 and uT16 in an unbound CPD-containing DNA duplex, Mu and co-workers in the Broyde laboratory estimated the flipping barriers via the major and minor grooves to be 9.6 and 10.5 kcal/mol, respectively, for the conformational capture mechanism, which can be compared with the present work. The observed difference in flipping barrier could be attributed to the fact that our model consisted on only 3T mismatch, while their contained the CPD lesion. However, Mu et al. 53 results for the correlated motion mechanism cannot be compared because this

13



part of their study entailed the entire binding free energy profile for RAD4 binding to the damaged duplex and not just base flipping. In general, the calculated activation barriers for extrusion of mismatched bases in BDI are in reasonable agreement with the values reported in the literature (Table S1 - Supporting Information) for DNA with different sequences and compositions.

Base Flipping Energetics in the Presence of RAD4 The binding of RAD4 to the mismatched DNA significantly alters the conformational free energy profiles of uT15 and uT16 with respect to those of the RAD4-free DNA. FREC (Φ) exhibits a broader energy well with a global free energy minimum located in the extra-helical region at Φ ∼-100o with an activation barrier of ∼11 kcal/mol at Φ ∼70o . Although the extrusion barrier height is the same for both the major- and minor-groove pathways, FREC (Φ) is relatively more shallow along the major-groove pathway than that along the minor-groove pathway (Figure S7 and Figure S8 - Supporting Information). The observed difference in FREC (Φ) between the major- and minor-groove pathways suggests that the major-groove pathway is the most likely path for the extra-helical intra-helical conformational transitions of uT16 in RAD4-DNA complex. FREC (Ψ) also exhibits a broader energy well with a global energy minimum in the extra-helical region at Ψ ∼123o (i.e., Ψmin =123o ) with an energy barrier of ∼15.3 kcal/mol observed in the intra-helical conformational region at Ψ ∼-76o . The observed global minimum and the activation barrier in FREC (Ψ) indicate that uT15 also prefers the extra-helical conformation in DNA-RAD4 complex.

RAD4 Binding-Induced Modulations of Conformational Energy Landscapes of Mismatched Bases FREC (Φ) versus FBDI (Φ) and FREC (Ψ) versus FBDI (Ψ) comparisons enable us to probe RAD4-induced changes in the conformational energy landscapes and flipping energetics of uT16 and uT15 bases of the mismatched DNA. Such a comparison reveals that the global free

14


Page 14 of 40

Page 15 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


energy minimum on the conformational landscape of uT16 is shifted from Φ ∼ 14o (in the absence of RAD4) to Φ ∼ -100o (in the presence of RAD4) upon RAD4 binding to DNA, and that the activation barrier for the flipping of uT16 is decreased from 12.5 kcal/mol in RAD4free DNA to 11 kcal/mol in RAD4-DNA complex. The observed RAD4-induced intra- to extra-helical conformational shift of uT16 and the overall reduction of the activation barrier separating the intra- and extra-helical states of uT16 may be essential for the catalytic role of RAD4 and for the stabilization of the extra-helical conformation of uT16 in RAD4-DNA complex (Figures S9-S12 - Supporting Information). The X-ray crystallographic structure-based RAD4-DNA binding mechanism suggested that insertion of the β-hairpin (VAL594 to PRO607) of BHD3 domain of RAD4 into the DNA duplex at the lesion site is an important driving force for the extrusion of uT16 and uT15 bases into the binding pocket of RAD4. 42 We estimated the energy penalty for the insertion of β-hairpin into the DNA to dislocate uT16 by calculating the observed increase in relative free energy at Φ=14o [FREC (Φ=14o )-FBDI (Φ=14o )] to be ∼7.5 kcal/mol. In addition, the reduction of free energy at Φ=-100o [FBDI (Φ=-100o )-FREC (Φ=-100o )], which was estimated to be ∼10.8 kcal/mol, quantifies the stabilizing energy provided by RAD4 to stabilize the extruded uT16 base in its extra-helical conformation.

A comparison of FREC (Ψ) and FBDI (Ψ) reveals that uT15 also exhibits a similar intrahelical→extra-helical conformational shift and barrier reduction upon binding of RAD4 to the mismatched DNA. Here again, the RAD4-induced increase in relative free energy at Ψ=37o [FREC (Ψ=37o )-FBDI (Ψ=37o )] of ∼8.5 kcal/mol provides a reasonable estimate of the energy cost for the insertion of the β-hairpin of RAD4 into the DNA duplex to dislocate uT15, while a decrease of ∼8.2 kcal/mol in free energy at Ψ=123o [FBDI (Ψ=123o )-FREC (Ψ=123o ] quantifies the stabilizing energy provided by RAD4 to stabilize uT15 in its extra-helical conformation.

15



Figure 5: The atomic structure of DNA-RAD4 complex in the highest-energy (Ψ ∼-76o , Φ ∼70o ) and lowest-energy (Ψ ∼123o , Φ ∼-100o ) conformational states of uT15 and uT16 bases (B: lowest-energy state of uT16; D: highest-energy state of uT16; F: lowest-energy state of uT15; H: highest-energy state of uT15). The corresponding NUCPLOTs of DNA-RAD4 interaction maps (A, C, E, G) showing critical interactions around the lesion are shown. The key residues of RAD4 (shown in CPK representation) that contribute to the conformational stability of uT15 and uT16 are color coded based on their strength of interactions with these mismatched bases. DNA-RAD4 Interactions and Conformational Stability of Mismatched Bases In order to quantitatively assess the important interactions stabilizing the minimum and transition states of the mismatched uT15 and uT16 bases in RAD4-DNA complex, the pair-

16


Page 16 of 40

Page 17 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


wise interactions between these mismatched bases and their neighboring residues of RAD4 were examined in the highest-(Ψ ∼-76o , Φ ∼70o ) and lowest-energy (Ψ ∼123o , Φ ∼-100o ) conformational states of uT15 and uT16. The pairwise interaction energies in the transition states were calculated from the umbrella sampling trajectories generated in windows centered at Ψ and Φ values corresponding to the transition states of RAD4-DNA complex. The corresponding pairwise energies at the most-stable conformations were calculated from trajectories obtained from direct MD simulations. The time-averaged structures obtained from the last 100 ps of the respective trajectories of the most-stable and transition states of RAD4DNA complex and their corresponding interaction maps (obtained using NUCPLOT 86 ) are shown in Figure 5. The distributions of the calculated energies of critical interactions between uT15, uT16, and important residues of RAD4 in the most-stable and transition states of RAD4-DNA complex are shown in Figure 6. The corresponding mean interaction energies of uT15 and uT16 are provided in Table 1. The results indicate that higher relative stability of the extrahelical conformations of uT15 and uT16 in RAD-DNA complex is primarily due to their interactions with BHD2 (ASP489 to TYR540) and BHD3 (TYR540 to ILE632) domains of RAD4.

17



0.2

1

1 uT15-PHE597

uT15-ARG494

uT15-MET498

0.15 0.1

0.5

0.5

P (E)

0.05 0 -30 -25 -20 -15 -10 -5

0

0 -10

-8

-6

-4

-2

0

0.4

1

0 -7 -6 -5 -4 -3 -2 -1 0 1.5

uT15-PHE599

uT15-ARG515

uT15-VAL605 1

0.2

0.5

0.5

0 -5

-4

-3

-2

-1

0

0 -20

-15

-10

-5

0

0 -4

-3

-2

-1

0

E (kcal/mol) 0.2

P (E )

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 40

1 0.8 uT16-ASN558 uT16-PHE556 0.15 uT16-ARG494 0.4 0.6 0.1 0.4 0.2 0.05 0.2 0 0 0 -8 -6 -4 -2 0 -30 -25 -20 -15 -10 -5 0 -10 -8 -6 -4 -2 1 1 uT16-PRO607 0.8 uT16-PHE597 0.5 0.8 uT16-ASN554 0.4 0.6 0.6 0.3 0.4 0.4 0.2 0.2 0.2 0.1 0 0 0 -7 -6 -5 -4 -3 -2 -1 0 -6 -5 -4 -3 -2 -1 0 -6 -5 -4 -3 -2 -1 1 1 2 uT16-LYS606 uT16-VAL594 0.8 0.8 uT16-VAL605 1.5 0.6 0.6 1 0.4 0.4 0.5 0.2 0.2 0 0 0 -2 -1 -5 -4 -3 -2 -1 0 -5 -4 -3 -2 -1 0

0

0

0

E (kcal/mol) Figure 6: Distributions of pairwise interaction energies between the mismatched bases of DNA and the key residues of RAD4 calculated in the most-stable (blue) and transition states (red) of the DNA-RAD4 complex are shown.

18


Page 19 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Role of ARG494 and ARG515 In particular, the calculated interaction energies reveal that ARG494 of BHD2 plays an important role in stabilizing uT15 and uT16 in their extra-helical conformations (mean interaction energies of uT15-ARG494 and uT16-ARG494 pairs are -16.67±7.7 and -6.66±7.3 kcal/mol, respectively). The strong ARG494-base interactions are due to the formation of hydrogen bonds between ARG494 and the phosphate groups of uT15 and uT16. In the minimum-energy extra-helical conformational state, the calculated distribution of uT15-ARG494 interaction energy exhibits two peaks at around -22.0 kcal/mol and 5.0 kcal/mol. The peak at -22.0 kcal/mol is due to configurations in which one or more hydrogen bonds are formed between the guanidinium (Gdn494) moiety of ARG494 and the phosphate (P15) unit of uT15. Similarly, the H-bonds between uT16 and ARG494 contribute to the peaks at -20 kcal/mol and -15 kcal/mol in the distribution of uT16-ARG494 interaction energy. ARG494 was observed to form one, two, and three hydrogen bonds with the mismatched partner bases in approximately 43%, 46%, and 3% of the configurations in the trajectory, respectively, while no H-bonds were observed in the remaining 8% of the configurations. Different types of more probable hydrogen bonds and their frequencies of occurrence in the MD trajectory are shown in Figure 7. In the most probable doubly hydrogen-bonded geometry (referred to as type A), a single H-bond is formed between an NH2 of ARG494 and a phosphate oxygen of uT15 and the other H-bond is formed between another NH2 of ARG494 and the glycosidic oxygen (O50 ) of uT15. In type B, two NH2 groups of ARG494 make two hydrogen bonds with two anionic phosphate oxygens of uT15, whereas in type C, a single phosphate oxygen of uT16 makes two hydrogen bonds with two NH2 groups of ARG494. In the most probable singly hydrogenbonded structures, an NH2 of ARG494 forms a hydrogen bond with a phosphate oxygen of uT15 or uT16 (as shown in Figure 7 D and E). The most probable triply hydrogen-bonded geometry is shown in Figure 7 F. The other metastable hydrogen-bonded configurations that are observed less frequently during the course of the simulation are also shown in Figure S27 19



(Supporting Information). The local structural rearrangements in the vicinity of the mismatched bases facilitate ARG494 to form H-bonds alternatively with uT15 and uT16 of the mismatched DNA. The dynamic nature of these H-bonds enables ARG494 to form H-bonds with either uT15 or uT16 for most of the time and, occasionally, to share its H-bonds between uT15 and uT16 during the course of the simulation (Figure S28). The configurations in which there are no H-bonds between uT15 and ARG494 contribute to the peak at -5.0 kcal/mol in the distribution of uT15-ARG494 interaction energy; in such configurations, ARG494 partners with and makes H-bonds with uT16. When ARG494 engages with uT16, uT15 occasionally forms H-bonds with ARG515 (for t < 2.5 ns); but the strength of uT15-ARG515 interaction is relatively less than that between uT15 and ARG494. The uT15-ARG494 interaction energy drops to -22.0 kcal/mol when uT15 reestablishes its H-bonds with ARG494. The observed coupled dynamics of uT15, uT16, ARG494, and ARG515 may be necessary for the balance of conformational stability and flexibility of mismatches partner bases in the global minimum state of the DNA-RAD4 complex. The electrostatic and van der Waals interactions of ARG494 and ARG515 with uT15 and uT16 also play an important role in the stabilization of the transition states of these mismatched partner bases (Figure S17-Supporting Information).

20


Page 20 of 40

Page 21 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 7: Different types of hydrogen bonding patterns between ARG494 and the mismatched partner bases observed from MD trajectory. (A) Type A: a single H-bond is formed between an NH2 of ARG494 and the phosphate oxygen (P15) of uT15 and the other H-bond is formed between another NH2 of ARG494 and the glycosidic oxygen (O50 ) of uT15. (B) Type B: two NH2 groups of ARG494 make two hydrogen bonds with two phosphate oxygens of uT15. (C) Type C: a single phosphate oxygen of uT16 makes two hydrogen bonds with two NH2 groups of ARG494. (D) Type D: ARG494 forms a single H-bond with a phosphate oxygen of uT15. (E) Type E: ARG494 forms a single H-bond with a phosphate oxygen of uT16. (F) Type F: a triply hydrogen-bonded geometry in which ARG494 forms H-bonds with phosphate oxygens of both uT15 and uT16.

In addition to ARG494, the non-covalent interactions of uT16 with PHE556, ASN558, ASN554, PRO607, PHE597, and VAL605 of BHD3 contribute significantly to the stability of the extra-helical conformation of uT16. Similarly, in the extra-helical state, uT15 experiences strong non-covalent interactions with PHE597, PHE599 and PHE562 of BHD3 and MET498, 21



ARG515, GLN495 of BHD2 domains of RAD4.

Role of Phenylalanine Residues Visual inspection of MD trajectories reveals that a set of four key phenylalanine residues (PHE556, PHE562, PHE597, and PHE599) of RAD4 favorably positioned near the lesion provide an aromatic pathway (also known as the phenyl flipping pathway 53 ) for extrusion of the orphaned bases in RAD4-DNA complex (see Figure S3 and Figure S4 - Supporting Information). The non-covalent interactions of PHE597, PHE599, PHE556, and PHE562 of RAD4 with uT15 and uT16 also contribute to the stability of the extra-helical conformation of RAD4-DNA complex. In particular, the pi-pi stacking (face-to-face) and T-shaped (edge-to-face) interactions between the phenyl rings of these phenylalanine residues and the pyrimidine rings of uT15 and uT16, and the sugar-pi interactions between the ribose sugars of uT15 and uT16 and the phenyl rings of these phenylalanine residues are critical for the extra-helical conformational stability of the mismatched partner bases. The calculated phenylalanine-base interaction energies also indicate that the non-covalent interactions between these phenylalanine residues and the mismatched nucleotide bases are crucial for the stabilization of the intermediate and the extra-helical conformational states of uT15 and uT16 during their extrusion via the aromatic pathway. In particular, uT16 interacts strongly with PHE556 and PHE597 of BHD3 domain of RAD4 by sandwiching between the two phenyl rings of these residues to optimize the stacking interactions between them. Similarly, uT15 is stabilized by its stacking interactions with PHE597 and PHE599 of BHD3 domain. uT15 interacts more strongly with PHE597 and PHE562 than it does with PHE599 and PHE556. In fact, uT15-PHE556 interaction is negligible compared to uT15-PHE597, ut15PHE562, and uT15-PHE599 interactions, which can be attributed to the fact that PHE556 is located relatively far from uT15 than other phenylalanine residues in RAD4-DNA complex. The calculated ranges of uT15-PHE interaction energies are as follows: uT15-PHE597 (from

22


Page 22 of 40

Page 23 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


-10 kcal/mol to -2 kcal/mol), uT15-PHE562 (from -7.57 kcal/mol to -0.08 kcal/mol), uT15PHE599 (from -4.14 kcal/mol to 0.98 kcal/mol), and uT15-PHE556 (fom -0.12 kcal/mol to 0.00 kcal/mol). In order to further characterize the nature of phenylalanine-base interactions in RAD4DNA complex, we examined the time evolution of the distances and the relative orientational angles between the phenylalanine residues and the mismatched partner bases. We define db and ds as the distances of the center of mass of the phenyl ring of a phenylalanine residue from the centers of mass of the pyrimidine ring and sugar ring of a mismatched partner base, respectively. Similarly, ηb and ηs denote the angles that the normal to the phenyl ring of a phenylalanine makes with the normal to the base ring and the sugar ring of a mismatched partner base, respectively. The suffices b and s denote the base and sugar of a mismatched partner nucleotide, respectively (Figure S18-Supporting Information). Figure 9 and Figures S19-S21 (Supporting Information) show the time evolution of interaction energy (E), db , ds , ηb , and ηs for all phenylalanine-base pairs. For uT15-PHE597 pair, db fluctuates around two different values (6.22 ˚ A and 4.70 ˚ A) and exhibits abrupt transitions between them at different time intervals (Figure 9). In particular, db fluctuates around 6.22 ˚ A (referred to as T-state) for t < 7.4 ns and 11 ns ≤ t ≤ 15.35 ns, while db fluctuates around 4.70 ˚ A (referred to as P-state) at all other times. The abrupt transitions between these two states observed at t ∼ 7.5 ns, t ∼ 10.8 ns, and t ∼ 15.5 ns can be attributed to structural rearrangements of uT15 and PHE597 in the vicinity of the mismatch site of RAD4-DNA complex. The mean values of db in the T-state and the P-state are 6.22±0.29 ˚ A and 4.70±0.35 ˚ A, respectively, which indicates that uT15 and PHE597 are relatively more proximal to each other in the P-state than in the T-state. The time evolution of uT15-PHE597 interaction energy (Figure 9) also shows similar transitions between -7.85 kcal/mol (corresponding to the P-state) and -5.25 kcal/mol (corresponding to the T-state) precisely at times coinciding with the transitions observed in the time evolution of db . The strength of uT15-PHE597 interaction is higher in the P-state

23



(mean energy is -7.85 kcal/mol) than that in the T-state (mean energy is -5.25 kcal/mol), which indicates that the P-state is relatively more stable than the T-state. The existence of P- and T-states provides a rationale for the observed bimodal distribution of uT15-PHE597 interaction energy; the first peak at -7.85 kcal/mol of this distribution corresponds to the P-state, while the second peak at -5.25 kcal/mol corresponds to the T-state.

Figure 8: Stacking geometries of the mismatched partner bases (uT15 and uT16) with PHE597 observed from MD trajectory. (A) Parallel-displaced uT15-PHE597 pi-pi stacking (face-to-face). The T-shaped stacking (edge-to-face) between (B) PHE597 and uT15 and (C and D) uT16 and PHE597. The time evolution of ηb sheds additional light on the structural changes accompanying the transitions between the P-state and T-state. Since ηb is a measure of the relative orientation of the phenyl ring of PHE597 and the pyrimidine ring of uT15, it quantifies the degree of stacking between these two moieties; ηb =0o for a perfectly stacked state, while ηb =90o for an ideal T-shaped stacking geometry. For uT15-PHE597, ηb fluctuates around 103.23o (with a mean of 103.23o ±12.95o ) in the T-state, while it fluctuates around 22.60o (with a mean of 22.60o ±11.34o ) in the P-state with similar transitions between these states as seen in the time evolution of the interaction energy and db . Based on the mean values 24


Page 24 of 40

Page 25 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


of db , ηb , and visual inspection of MD trajectories, it is inferred that, in the P-state, the phenyl ring of PHE597 forms a distorted parallel-displaced stacked geometry (as shown in Figure 8 A) with uT15, while, in the T-state, they adopt a distorted T-shaped geometry (as shown in Figure 8 B). The time evolution of interaction energy, db , ds , ηb , and ηs calculated for the PHE597-uT16 pair did not show any noticeable transitions during the course of the simulation. The calculated mean of uT16-PHE597 interaction energy is -3.62±0.84 kcal/mol (Figure 8.C, D)), which is less than that for uT15-PHE597 pair, which indicates the interaction strength of uT16-PHE597 is relatively less than that of uT15-PHE597 in RAD4-DNA complex. The mean values of db and ds are 6.09±0.33 ˚ A and 5.96±0.29 ˚ A, respectively for uT16-PHE597 (Figure 9). ηb fluctuates around 70.76o (with a mean of 70.76o ±9.66o ), while ηs exhibits transitions between 111.61o (for T-state; the mean value of ηs is 111.61o ± 14.12o ) and 135.61o (for P-state; the mean is 135.61o ±9.58o ), respectively. These transitions coincide with those observed for the uT15-PHE597 pair and they are due to slight reorientation of the sugar of uT16 in RAD4-DNA complex. To characterize the proximity and relative orientation of the sugar ring of uT15 with respect to PHE597, the time evolution of ds and ηs was examined (Figure 9). ηs also exhibits transitions coinciding with the transitions observed in the time evolution of db and ηb , but the degree of orientational change associated with the transitions in ηs is relatively less than that observed in ηb . The mean values of ηs in the T-state and the P-state are 156.64o ±9.0o A ±0.36 ˚ A with and 119.44o ±9.14o , respectively. ds fluctuates around a mean value of 4.4 ˚ no abrupt transitions. A similar bimodal distribution of interaction energy was observed for uT15-PHE562 pair. Like PHE597, PHE562 also forms two different packing geometries with uT15; The mean values of interaction energy, db , and ηb , in the most stable state are -5.25±1.11 kcal/mol, 4.86±0.35 ˚ A, and 70.39o ±13.30o , respectively (Figure S20 - Supporting Information). In this state, PHE562 forms a distorted T-shaped pi-pi stacking geometry with the base of uT15. The corresponding values in the alternate state are -0.66±0.56 kcal/mol, 7.27 ˚ A ±0.65 ˚ A,

25



and 123.16o ±16.28o . Since the mean distance of sugar of uT15 from the phenyl ring of PHE52 is greater than 7 ˚ A in both states, sugar-pi interaction is relatively weaker than the pi-pi interaction between uT15 and PHE562. The results also show that uT16-PHE562 interaction strength is negligibly small in RAD4-DNA complex. PHE599 interacts primarily with uT15 and the calculated mean of uT15-PHE599 interaction energy is -2.34±0.67 kcal/mol (Figure S23 -Supporting Information). No noticeable transitions were observed in the time evolution of uT15-PHE599 interaction energy. The calculated mean values of the distance and angle parameters for uT15-PHE599 pair are db =6.83 ˚ A ±0.57 ˚ A, ds =5.94 ˚ A ±0.42 ˚ A, ηb =91.92o ±37.65o and ηs =148.13o ±12.24o (Figure S21 -Supporting Information). PHE556 interacts primarily with uT16 with a mean interaction energy of -7.16±0.86 kcal/mol (Figure S22 -Supporting Information). The mean values of the distance and angle parameters for uT16-PHE556 are db =3.77 ˚ A± 0.27 ˚ A, ds =5.52 ˚ A± 0.28 ˚ A, ηb =162.49o ±8.50o and ηs =107.52o ±13.24o (Figure S19 -Supporting Information). The stronger uT16-PHE556 interaction is due to proximity of PHE556 to uT16 and the parallel alignment of the phenyl ring of PHE556 and the pyrimidine ring of uT16.

26


Page 26 of 40

Page 27 of 40

0

100 0

5

10

15

20

0

Time (ns)

5

10

15

b

150 100

s

b

η (°)

s

150

4 150 100 50 0

η (°)

6

b

η (°)

ds(Å)

db(Å)

7 6 5 8

d s(Å)

-8 -12 8 6 4 6 5 4 3 150 100 50 0

d (Å)

2 0 -2 E -4 -6

E -4

η (°)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


20

Figure 9: Time evolution of the interaction energy (E in kcal/mol) and the distance (db and ds ) and angle (ηb and ηs ) parameters for uT15-PHE597 (left panels) and uT16-PHE597 (right panels) pairs in RAD4-DNA complex.

The calculated mean interaction energy between MET498 and uT15 in the extra-helical state is -4.59±1.16 kcal/mol, of which -4.42±1.04 kcal/mol is due to van der waals interactions between them (Figure S26-Supporting Information). In the transition state, the conformational changes in uT15 increase the distance between uT15 and MET498, which in turn results in weakening of the van der Waals interactions between them (Figure S24 -Supporting Information). ASN554 and ASN558 form H-bonds with the NH2 groups of ASN554 and ASN558 point toward the O4 of uT16 to from H-bonds with it (Figure S25 -Supporting Information). The results obtained from the interaction energy-based analyses not only enabled us to identify key residues contributing to the conformational stability of the mismatched bases but also provided quantitative estimates of their relative energetic contributions to the stability of uT15 and uT16 bases in RAD4-DNA complex.

27



Table 1: Mean base-residue interaction energies of the mismatched bases (uT15 and uT16) min with the key residues of RAD4 in the most-stable conformational state (hEimin uT 15 and hEiuT 16 ) TS TS and the transition state (hEiuT 15 and hEiuT 16 ) of RAD4-DNA complex. Residues ARG494 GLN495 MET498 ARG515 ASN554 PHE556 ASN558 PHE562 VAL594 PHE597 PHE599 VAL605 LYS606 PRO607

TS min TS hEimin uT 15 (kcal/mol) hEiuT 15 (kcal/mol) hEiuT 16 (kcal/mol) hEiuT 16 (kcal/mol) -16.67±7.70 -11.31±4.90 -6.66±7.30 -13.36±5.32 -1.73±1.79 -0.16±0.16 -0.07±0.07 -4.59±1.16 -0.29±0.36 -0.26±0.22 -0.27±0.47 -1.82±4.66 -5.86±4.58 -0.01±0.08 -0.48±0.28 -4.81±1.18 -0.04±0.01 -0.03±0.02 -7.16±0.86 -0.11±0.31 -6.21±0.89 -2.47±2.38 0.05±0.04 -0.22±0.12 -0.11±0.10 -1.42±0.32 -6.16±1.55 -0.99±1.07 -3.62±0.84 -0.36±0.28 -2.33±0.66 -0.30±0.38 -0.11±0.05 -1.29±0.51 -0.57 ±0.62 -2.23±0.06 -0.31±0.26 -0.16±0.08 -0.02±0.03 -1.63±0.83 -0.02±0.08 -0.15±0.08 -0.02±0.02 -3.71±0.67 -0.02±0.12

Conclusions In summary, the present study examined the molecular mechanism, dynamics, and energetics of RAD4-mediated dinucleotide base extrusion in a mismatched DNA (with three contiguous thymidine-thymidine mismatches) using molecular dynamics simulation and umbrella sampling method. The mismatched uT15 and uT16 bases adopted the most stable intra-helical conformation and their extrusion was unfavorable in RAD4-free mismatched DNA due to the presence of prohibitively high barriers (> 12 kcal/mol) along the extrusion pathways. The results indicate that uT15 is relatively more flexible than uT16 in the intra-helical conformation of RAD4-free DNA, and that both uT15 and uT16 are likely to prefer their respective major groove pathways for their extrusion.The binding of RAD4 to DNA induces intra-helical → extra-helical conformational transitions in uT15 and uT16 indicating the key role of RAD4-DNA interactions in base extrusion and extra-helical stabilization of these mismatched bases in the RAD4-DNA complex. The calculated energy penalties for insertion of β-hairpin of RAD4 into DNA duplex to dislocate uT15 and uT16 are 8.5 kcal/mol and 7.5 kcal/mol, respectively and the calculated extra-helical stabilization energies of uT15 and

28


Page 28 of 40

Page 29 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


uT16 in RAD4-DNA complex are 8.2 kcal/mol and 10.8 kcal/mol, respectively. A detailed examination of critical RAD4-DNA interactions reveals that ARG494, PHE556, ASN558, ASN554, VAL594, PHE597, PHE559, ARG515, GLN495, MET498, VAL605, and PRO607 of RAD4 are key residues contributing to the conformational stability of uT15 and uT16 in RAD4-DNA complex. The observed RAD4-induced conformational shift of mismatched bases support the correlated motion mechanism for the RAD4-mediated DNA damage repair process.

Acknowledgement We wish to thank Dr. Moumita Saharay of Osmania University for critical reading of the manuscript and Dr. A. Semparithi for computational assistance and support. The highperformance computing facility provided by IIIT-Hyderabad is gratefully acknowledged. K.P. acknowledges CSIR, Govt. of India, for a senior research fellowship

References (1) Xue, W.; Warshawsky, D. Metabolic Activation of Polycyclic and Heterocyclic Aromatic Hydrocarbons and DNA Damage: A Review. Toxicol. Appl. Pharmacol. 2005, 206, 73– 93. (2) Sproviero, M.; Verwey, A. M.; Rankin, K. M.; Witham, A. A.; Soldatov, D. V.; Manderville, R. A.; Fekry, M. I.; Sturla, S. J.; Sharma, P.; Wetmore, S. D. Structural and Biochemical Impact of C8-aryl-guanine Adducts Within the Nar I Recognition DNA Sequence: Influence of Aryl Ring size on Targeted and Semi-targeted Mutagenicity. Nucleic Acids res. 2014, 42, 13405–13421. (3) Mukherjee, A.; Lavery, R.; Bagchi, B.; Hynes, J. T. On the Molecular Mechanism of

29



Drug Intercalation into DNA: A Simulation Study of the Intercalation Pathway, Free Energy, and DNA Structural Changes. J. Am. Chem. Soc. 2008, 130, 9747–9755. (4) Helleday, T.; Eshtad, S.; Nik-Zainal, S. Mechanisms Underlying Mutational Signatures in Human Cancers. Nat. Rev. Genet. 2014, 15, 585–598. (5) Lindahl, T. Instability and Decay of the Primary Structure of DNA. Nature 1993, 362, 709–715. (6) Golub, T. R.; Slonim, D. K.; Tamayo, P.; Huard, C.; Gaasenbeek, M.; Mesirov, J. P.; Coller, H.; Loh, M. L.; Downing, J. R.; Caligiuri, M. A. Molecular Classification of Cancer: Class Discovery and Class Prediction by Gene Expression Monitoring. Science 1999, 286, 531–537. (7) Van’t Veer, L. J.; Dai, H.; Van De Vijver, M. J.; He, Y. D.; Hart, A. A.; Mao, M.; Peterse, H. L.; Van Der Kooy, K.; Marton, M. J.; Witteveen, A. T. Gene Expression Profiling Predicts Clinical Outcome of Breast Cancer. Nature 2002, 415, 530–536. ¨ (8) Sancar, A.; Lindsey-Boltz, L. A.; Unsal-Ka¸ cmaz, K.; Linn, S. Molecular Mechanisms of Mammalian DNA Repair and the DNA Damage Checkpoints. Annu. Rev. Biochem. 2004, 73, 39–85. (9) Stein, B.; Rahmsdorf, H. J.; Steffen, A.; Litfin, M.; Herrlich, P. UV-induced DNA Damage is an Intermediate Step in UV-induced Expression of Human Immunodeficiency Virus Type-1, Collagenase, C-fos, and Metallothionein. Mol. Cell. Biol. 1989, 9, 5169– 5181. (10) Dizdaroglu, M. Chemical Determination of Free Radical-induced Damage to DNA. Free Radic. Biol. Med. 1991, 10, 225–242. (11) Singh, K. K. The Saccharomyces Cerevisiae Sln1p-Ssk1p Two-component System Me-

30


Page 30 of 40

Page 31 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


diates Response to Oxidative Stress and in an Oxidant-specific Fashion. Free Radic. Biol. Med. 2000, 29, 1043–1050. (12) Mitchell, J. R.; Hoeijmakers, J. H.; Niedernhofer, L. J. Divide and Conquer: Nucleotide Excision Repair Battles Cancer and Ageing. Curr. Opin. Struct. Biol. 2003, 15, 232– 240. (13) Strand, M.; Prolla, T. A.; Liskay, R. M.; Petes, T. D. Destabilization of Tracts of Simple Repetitive DNA in Yeast by Mutations Affecting DNA Mismatch Repair. Nature 1993, 365, 274–276. (14) D’Errico, M.; Teson, M.; Calcagnile, A.; De Santis, L. P.; Nikaido, O.; Botta, E.; Zambruno, G.; Stefanini, M.; Dogliotti, E. Apoptosis and Efficient Repair of DNA Damage Protect Human Keratinocytes Against UVB. Cell Death Differ. 2003, 10 . (15) Kuraoka, I.; Bender, C.; Romieu, A.; Cadet, J.; Wood, R. D.; Lindahl, T. Removal of Oxygen Free-radical-induced 5, 8-purine Cyclodeoxynucleosides from DNA by the Nucleotide Excision-Repair Pathway in Human Cells. Proc. Natl. Acad. Sci. 2000, 97, 3832–3837. (16) Gillet, L. C.; Schärer, O. D. Molecular Mechanisms of Mammalian Global Genome Nucleotide Excision Repair. Chem. Rev. 2006, 106, 253–276. (17) Wang, X. W.; Yeh, H.; Schaeffer, L.; Roy, R.; Moncollin, V.; Egly, J.-M.; Wang, Z.; Friedberg, E.; Evans, M.; Taffe, B. P53 Modulation of TFIIH-associated Nucleotide Excision Repair Activity. Nat. Genet. 1995, 10, 188–195. (18) Evans, E.; Moggs, J. G.; Hwang, J. R.; Egly, J.-M.; Wood, R. D. Mechanism of Open Complex and Dual Incision Formation by Human Nucleotide Excision Repair Factors. EMBO J. 1997, 16, 6559–6573.

31



(19) Liaw, D.; Marsh, D. J.; Li, J.; Dahia, P. L.; Wang, S. I.; Zheng, Z.; Bose, S.; Call, K. M.; Tsou, H. C.; Peacoke, M. Germline Mutations of the PTEN Gene in Cowden Disease, an Inherited Breast and Thyroid Cancer Syndrome. Nat. Genet. 1997, 16, 64–67. (20) Botstein, D.; Risch, N. Discovering Genotypes Underlying Human Phenotypes: Past Successes for Mendelian Disease, Future Approaches for Complex Disease. Nat. Genet. 2003, 33, 228. (21) Jackson, S. P.; Bartek, J. The DNA-damage Response in Human Biology and Disease. Nature 2009, 461, 1071. (22) Parikh, S. S.; Mol, C. D.; Slupphaug, G.; Bharati, S.; Krokan, H. E.; Tainer, J. A. Base Excision Repair Initiation Revealed by Crystal Structures and Binding Kinetics of Human Uracil-DNA Glycosylase with DNA. EMBO J. 1998, 17, 5214–5226. (23) Jamieson, E. R.; Lippard, S. J. Structure, Recognition, and Processing of CisplatinDNA Adducts. Chem. Rev. 1999, 99, 2467–2498. (24) Chen, X.; Velmurugu, Y.; Zheng, G.; Park, B.; Shim, Y.; Kim, Y.; Liu, L.; Van Houten, B.; He, C.; Ansari, A. Kinetic Gating Mechanism of DNA Damage Recognition by Rad4/XPC. Nat. Commun. 2015, 6, 5849. (25) Sinha, R. P.; Häder, D.-P. UV-induced DNA Damage and Repair: A Review. Photochem. Photobiol. Sci. 2002, 1, 225–236. (26) Burns, J. L.; Guzder, S. N.; Sung, P.; Prakash, S.; Prakash, L. An Affinity of Human Replication Protein A for Ultraviolet-damaged DNA Implications for Damage Recognition in Nucleotide Excision Repair. J. Biol. Chem. 1996, 271, 11607–11610. (27) Jansen, L. E.; Verhage, R. A.; Brouwer, J. Preferential Binding of Yeast Rad4· Rad23 Complex to Damaged DNA. J. Biol. Chem. 1998, 273, 33111–33114.

32


Page 32 of 40

Page 33 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(28) Batty, D. P.; Wood, R. D. Damage Recognition in Nucleotide Excision Repair of DNA. Gene 2000, 241, 193–204. (29) Araki, M.; Masutani, C.; Takemura, M.; Uchida, A.; Sugasawa, K.; Kondoh, J.; Ohkuma, Y.; Hanaoka, F. Centrosome Protein Centrin 2/caltractin 1 is Part of the Xeroderma Pigmentosum group C Complex that Initiates Global Genome Nucleotide Excision Repair. J. Biol. Chem. 2001, 276, 18665–18672. (30) Nishi, R.; Okuda, Y.; Watanabe, E.; Mori, T.; Iwai, S.; Masutani, C.; Sugasawa, K.; Hanaoka, F. Centrin 2 Stimulates Nucleotide Excision Repair by Interacting with Xeroderma Pigmentosum Group C Protein. Mol.Cell. Biol. 2005, 25, 5664–5674. (31) Fujiwara, Y.; Masutani, C.; Mizukoshi, T.; Kondo, J.; Hanaoka, F.; Iwai, S. Characterization of DNA Recognition by the Human UV-damaged DNA-binding Protein. J. Biol. Chem. 1999, 274, 20027–20033. (32) Li, T.; Chen, X.; Garbutt, K. C.; Zhou, P.; Zheng, N. Structure of DDB1 in Complex with a Paramyxovirus V Protein: Viral Hijack of a Propeller Cluster in Ubiquitin Ligase. Cell. 2006, 124, 105–117. (33) Ghodke, H.; Wang, H.; Hsieh, C. L.; Woldemeskel, S.; Watkins, S. C.; Rapić-Otrin, V.; Van Houten, B. Single-molecule Analysis Reveals Human UV-damaged DNA-binding Protein (UV-DDB) Dimerizes on DNA via Multiple Kinetic Intermediates. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, E1862–E1871. (34) Sugasawa, K. Multiple DNA Damage Recognition Factors Involved in Mammalian Nucleotide Excision Repair. Biochemistry (Moscow) 2011, 76, 16–23. (35) Velmurugu, Y.; Chen, X.; Sevilla, P. S.; Min, J.-H.; Ansari, A. Twist-open Mechanism of DNA Damage Recognition by the Rad4/XPC Nucleotide Excision Repair Complex. Proc. Natl. Acad. Sci. 2016, 113, E2296–E2305.

33



(36) de Laat, W. L.; Jaspers, N. G.; Hoeijmakers, J. H. Molecular Mechanism of Nucleotide Excision Repair. Genes Dev. 1999, 13, 768–785. (37) Yokoi, M.; Masutani, C.; Maekawa, T.; Sugasawa, K.; Ohkuma, Y.; Hanaoka, F. The Xeroderma Pigmentosum group C Protein Complex XPC-HR23B Plays an Important Role in The Recruitment of Transcription Factor IIH to Damaged DNA. J. Biol. Chem. 2000, 275, 9870–9875. (38) Sugasawa, K.; Okamoto, T.; Shimizu, Y.; Masutani, C.; Iwai, S.; Hanaoka, F. A Multistep Damage Recognition Mechanism for Global Genomic Nucleotide Excision Repair. Genes Dev. 2001, 15, 507–521. (39) Sugasawa, K.; Shimizu, Y.; Iwai, S.; Hanaoka, F. A Molecular Mechanism for DNA Damage Recognition by the Xeroderma Pigmentosum group C Protein Complex. DNA Repair 2002, 1, 95–107. (40) Sugasawa, K. XPC: its product and biological roles. Mol. Mech. Xer. Pigmen. 2009, 47–56. (41) Kong, M.; Liu, L.; Chen, X.; Driscoll, K. I.; Mao, P.; Böhm, S.; Kad, N. M.; Watkins, S. C.; Bernstein, K. A.; Wyrick, J. J. Single-molecule Imaging Reveals that Rad4 Employs a Dynamic DNA Damage Recognition Process. Mol. Cell. 2016, 64, 376–387. (42) Min, J.-H.; Pavletich, N. P. Recognition of DNA Damage by the Rad4 Nucleotide Excision Repair Protein. Nature 2007, 449, 570–575. (43) Schärer, O. D. Nucleotide Excision Repair in Eukaryotes. Cold Spring Harb. Perspect. Biol. 2013, 5, a012609. (44) Sugasawa, K.; Akagi, J.-i.; Nishi, R.; Iwai, S.; Hanaoka, F. Two-step Recognition of

34


Page 34 of 40

Page 35 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


DNA Damage for Mammalian Nucleotide Excision Repair: Directional Binding of the XPC Complex and DNA Strand Scanning. Mol. Cell. 2009, 36, 642–653. (45) Camenisch, U.; Dip, R.; Schumacher, S. B.; Schuler, B.; Naegeli, H. Recognition of Helical Kinks by Xeroderma Pigmentosum Group A Protein Triggers DNA Excision Repair. Nat. Struct. Mol. Biol. 2006, 13, 278. (46) Baute, J.; Depicker, A. Base Excision Repair and its Role in Maintaining Genome Stability. Crit. Rev. Biochem. Mol. Biol. 2008, 43, 239–276. (47) Sakurai, S.; Kitano, K.; Yamaguchi, H.; Hamada, K.; Okada, K.; Fukuda, K.; Uchida, M.; Ohtsuka, E.; Morioka, H.; Hakoshima, T. Structural Basis for Recruitment of Human Flap Endonuclease 1 to PCNA. EMBO J. 2005, 24, 683–693. (48) Liu, Y.; Kao, H.-I.; Bambara, R. A. Flap Endonuclease 1: A Central Component of DNA Metabolism. Annu. Rev. Biochem. 2004, 73, 589–615. (49) Cassimeris, L.; Plopper, G.; Lingappa, V. R. Lewin’s Cells; Jones & Bartlett Publishers, 2011. (50) Park, S.-Y.; Lam, W.; Cheng, Y.-c. X-ray Repair Cross-complementing Gene I Protein Plays an Important Role in Camptothecin Resistance. Cancer Res. 2002, 62, 459–465. (51) Marteijn, J. A.; Lans, H.; Vermeulen, W.; Hoeijmakers, J. H. Understanding Nucleotide Excision Repair and its Roles in Cancer and Ageing. Nat. Rev. Mol. Cell Biol. 2014, 15, 465. (52) Bowman, G. D.; O’donnell, M.; Kuriyan, J. Structural Analysis of a Eukaryotic Sliding DNA Clamp-clamp Loader Complex. Nature 2004, 429, 724–730. (53) Mu, H.; Geacintov, N. E.; Zhang, Y.; Broyde, S. Recognition of Damaged DNA for Nucleotide Excision Repair: A Correlated Motion Mechanism with a Mismatched cissyn Thymine Dimer Lesion. Biochemistry 2015, 54, 5263–5267. 35



(54) Camenisch, U.; Träutlein, D.; Clement, F. C.; Fei, J.; Leitenstorfer, A.; FerrandoMay, E.; Naegeli, H. Two-stage Dynamic DNA Quality Check by Xeroderma Pigmentosum Group C Protein. The EMBO journal 2009, 28, 2387–2399. (55) Imhof, P.; Zahran, M. The Effect of a G:T Mispair on the Dynamics of DNA. PloS One 2013, 8, e53305. (56) Banavali, N. K.; MacKerell, A. D. Free Energy and Structural Pathways of Base Flipping in a DNA GCGC Containing Sequence. J. Mol. Biol. 2002, 319, 141–160. (57) Qi, Y.; Spong, M. C.; Nam, K.; Banerjee, A.; Jiralerspong, S.; Karplus, M.; Verdine, G. L. Encounter and Extrusion of an Intrahelical Lesion by a DNA Repair Enzyme. Nature 2009, 462, 762–766. (58) Wang, P.; Brank, A. S.; Banavali, N. K.; Nicklaus, M. C.; Marquez, V. E.; Christman, J. K.; MacKerell, A. D. Use of Oligodeoxyribonucleotides with Conformationally Constrained Abasic Sugar Targets to Probe the Mechanism of Base Flipping by Hha I DNA (Cytosine C5)-methyltransferase. J. Am. Chem. Soc. 2000, 122, 12422–12434. (59) Best, R. B.; Hummer, G. Reaction Coordinates and Rates from Transition Paths. Proc. Natl. Acad. Sci. 2005, 102, 6732–6737. (60) Bolhuis, P. G.; Chandler, D.; Dellago, C.; Geissler, P. L. Transition Path Sampling: Throwing Ropes Over Rough Mountain Passes, in the Dark. Annu. Rev. Phys. Chem. 2002, 53, 291–318. (61) Bartels, C.; Karplus, M. Probability Distributions for Complex Systems: Adaptive Umbrella Sampling of the Potential Energy. J. Phys. Chem. B. 1998, 102, 865–880. (62) Banerjee, A.; Yang, W.; Karplus, M.; Verdine, G. L. Structure of a Repair Enzyme Interrogating Undamaged DNA Elucidates Recognition of Damaged DNA. Nature 2005, 434, 612–618. 36


Page 36 of 40

Page 37 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(63) Mu, Y.; Nguyen, P. H.; Stock, G. Energy Landscape of a Small Peptide Revealed by Dihedral Angle Principal Component Analysis. Proteins Struct. Funct. Bioinforma. 2005, 58, 45–52. (64) Giudice, E.; Várnai, P.; Lavery, R. Basepair Opening within B-DNA: Free Energy Pathways for GC and AT Pairs from Umbrella Sampling Simulations. Nucleic Acids Res. 2003, 31, 1434–1443. (65) Song, K.; Campbell, A. J.; Bergonzo, C.; de los Santos, C.; Grollman, A. P.; Simmerling, C. An Improved Reaction Coordinate for Nucleicacid Base Flipping Studies. J. Chem. Theory Comput. 2009, 5, 3105–3113. (66) Parker, J. B.; Bianchet, M. A.; Krosky, D. J.; Friedman, J. I.; Amzel, L. M.; Stivers, J. T. Enzymatic Capture of an Extrahelical Thymine in the Search for Uracil in DNA. Nature 2007, 449, 433–437. (67) Cao, C.; Jiang, Y. L.; Stivers, J. T.; Song, F. Dynamic Opening of DNA During the Enzymatic Search for a Damaged Base. Nat. Struct. Mol. Biol. 2004, 11, 1230–1236. (68) Cai, Y.; Zheng, H.; Ding, S.; Kropachev, K.; Schwaid, A. G.; Tang, Y.; Mu, H.; Wang, S.; Geacintov, N. E.; Zhang, Y. Free Energy Profiles of Base Flipping in Intercalative Polycyclic Aromatic Hydrocarbon-damaged DNA Duplexes: Energetic and Structural Relationships to Nucleotide Excision Repair Susceptibility. Chem. Res. Toxicol. 2013, 26, 1115–1125. (69) Mu, H.; Geacintov, N. E.; Min, J.-H.; Zhang, Y.; Broyde, S. Nucleotide Excision Repair Lesion-recognition Protein Rad4 Captures a Pre-flipped Partner Base in a Benzo [a] pyrene-derived DNA lesion: How Structure Impacts the Binding Pathway. Chem. Res. Toxicol. 2017, (70) Webb, B.; Sali, A. Comparative Protein Structure Modeling Using Modeller. Curr. Protoc. Protein. Sci. 2014, 2–9. 37



(71) Humphrey, W.; Dalke, A.; Schulten, K. VMD: visual Molecular Dynamics. J. Mol. Graph. 1996, 14, 33–38. (72) Berendsen, H. J.; van der Spoel, D.; van Drunen, R. GROMACS: A Message-passing Parallel Molecular Dynamics Implementation. Comput. Phys. Commun. 1995, 91, 43– 56. (73) Hess, B.; Kutzner, C.; Van Der Spoel, D.; Lindahl, E. GROMACS 4: Algorithms for Highly Efficient, Load-balanced, and Scalable Molecular Simulation. J. Chem. Theory Comput. 2008, 4, 435–447. (74) Ivani, I.; Dans, P. D.; Noy, A.; Ramon,; Balaceanu, A.; Portella, G. Parmbsc1: A Refined Forcefield for DNA Simulations. Nature methods 2016, 13, 55–58. (75) Hess, B. P-LINCS: A Parallel Linear Constraint Solver for Molecular Simulation. J. Chem. Theory Comput. 2008, 4, 116–122. (76) Darden, T.; York, D.; Pedersen, L. Particle Mesh Ewald: An N-log (N) Method for Ewald Sums in Large Systems. J. Chem. Phys. 1993, 98, 10089–10092. (77) Essmann, U.; Perera, L.; Berkowitz, M. L.; Darden, T.; Lee, H.; Pedersen, L. G. A Smooth Particle Mesh Ewald Method. J. Chem. Phys. 1995, 103, 8577–8593. (78) Nosé, S.; Klein, M. Constant Pressure Molecular Dynamics for Molecular Systems. Mol. Phys. 1983, 50, 1055–1076. (79) Parrinello, M.; Rahman, A. Polymorphic Transitions in Single Crystals: A New Molecular Dynamics Method. J. Appl. Phys. 1981, 52, 7182–7190. (80) Bussi, G.; Donadio, D.; Parrinello, M. Canonical Sampling Through Velocity Rescaling. J. Chem. Phys. 2007, 126, 014101. (81) Torrie, G. M.; Valleau, J. P. Nonphysical Sampling Distributions in Monte Carlo Freeenergy Estimation: Umbrella Sampling. J. Comput. Phys. 1977, 23, 187–199. 38


Page 38 of 40

Page 39 of 40 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(82) Bonomi, M.; Branduardi, D.; Bussi, G.; Camilloni, C.; Provasi, D.; Raiteri, P.; Donadio, D.; Marinelli, F.; Pietrucci, F.; Broglia, R. A. PLUMED: A Portable Plugin for Free-energy Calculations with Molecular Dynamics. Comput. Phys. Commun. 2009, 180, 1961–1972. (83) Grossfield, A. An Implementation of WHAM: the Weighted Histogram Analysis Method Version 2.0. 9. Comput. Phys. Comm 1995, 91, 275–282. (84) Klimovich, P. V.; Shirts, M. R.; Mobley, D. L. Guidelines for the Analysis of Free Energy Calculations. J. Comput. Aided Mol. Des. 2015, 29, 397–411. (85) Rajitha R, J. C., Smith; Krishnan, M. Hidden Regularity and Universal Classification of Fast Side Chain Motions in Proteins. J. Am. Chem. Soc. 2014, 136, 8590–8605. (86) Luscombe, N. M.; Laskowski, R. A.; Thornton, J. M. NUCPLOT: A Program to Generate Schematic Diagrams of Protein-nucleicacid Interactions. Nucleic Acids Res. 1997, 25, 4940–4945.

39



Graphical TOC Entry

40


Page 40 of 40

Molecular Mechanism, Dynamics, and Energetics of Protein-Mediated

Recommend Documents