Molecular Mechanism underlying PRMT1 Dimerization for SAM

Nov 12, 2015 - Molecular Mechanism underlying PRMT1 Dimerization for SAM Binding and Methylase Activity. Ran Zhou†‡, Yiqian Xie‡, Hao Hu§, Guan...
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Molecular Mechanism underlying PRMT1 Dimerization for SAM Binding and Methylase Activity Ran Zhou,†,‡,⊥ Yiqian Xie,‡,⊥ Hao Hu,§,⊥ Guang Hu,† Viral Sanjay Patel,§ Jin Zhang,∥ Kunqian Yu,‡ Yiran Huang,∥ Hualiang Jiang,‡ Zhongjie Liang,*,† Yujun George Zheng,*,§ and Cheng Luo*,‡ †

Center for Systems Biology, Soochow University, Jiangsu 215006, China State Key Laboratory of Drug Research, Shanghai Institute of Materia Medica, Chinese Academy of Sciences, Shanghai 201203, China § Department of Pharmaceutical and Biomedical Sciences, University of Georgia, Athens, Georgia 30602, United States ∥ Department of Urology, Renji Hospital, School of Medicine, Shanghai Jiao Tong University, Shanghai 201203, China ‡

S Supporting Information *

ABSTRACT: Protein arginine methyltransferases (PRMTs) catalyze the posttranslational methylation of arginine, which is important in a range of biological processes, including epigenetic regulation, signal transduction, and cancer progression. Although previous studies of PRMT1 mutants suggest that the dimerization arm and the N-terminal region of PRMT1 are important for activity, the contributions of these regions to the structural architecture of the protein and its catalytic methylation activity remain elusive. Molecular dynamics (MD) simulations performed in this study showed that both the dimerization arm and the N-terminal region undergo conformational changes upon dimerization. Because a correlation was found between the two regions despite their physical distance, an allosteric pathway mechanism was proposed based on a network topological analysis. The mutation of residues along the allosteric pathways markedly reduced the methylation activity of PRMT1, which may be attributable to the destruction of dimer formation and accordingly reduced S-adenosyl-L-methionine (SAM) binding. This study provides the first demonstration of the use of a combination of MD simulations, network topological analysis, and biochemical assays for the exploration of allosteric regulation upon PRMT1 dimerization. These findings illuminate the results of mechanistic studies of PRMT1, which have revealed that dimer formation facilitates SAM binding and catalytic methylation, and provided direction for further allosteric studies of the PRMT family.

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INTRODUCTION Arginine methylation is a widespread post-translational modification in eukaryotic organisms and is catalyzed by protein arginine methyltransferases (PRMTs).1,2 This methylation results in epigenetic modifications of histones or changes in protein−protein interactions, which in turn lead to the regulation of a wide range of fundamental biological processes, including transcriptional activation/repression, signal transduction, chromatin remodeling, cell differentiation, and embryonic development.3,4 In recent years, the dysregulation of PRMTs was shown to be closely related to human diseases, particularly cancer and cardiovascular disease, and these enzymes have thus been considered to be potential therapeutic targets in clinical treatment.4−7 In methylation, PRMTs transfer the methyl group from Sadenosyl-L-methionine (SAM, also known as AdoMet) to the side-chain nitrogens of arginine residues, forming methylated arginine derivatives and S-adenosyl-L-homocysteine (SAH, also known as AdoHcy) as the products. To date, 11 PRMT isoforms have been identified,3 and in mammalian cells, nine © XXXX American Chemical Society

PRMTs (PRMT1 to PRMT9) have been classified into two main classes (type I and II).5 Both classes catalyze the formation of monomethyl arginine (MMA) but differ in the dimethyl arginine products obtained from the reaction: type I PRMTs (PRMT1-4, 6, and 8) catalyze asymmetric dimethyl arginine (ADMA) formation,8−12 whereas type II PRMTs (PRMT5 and 9) catalyze symmetric dimethyl arginine (SDMA) formation.13,14 PRMT7 was recently characterized as a type III PRMT because it only catalyzes the production of MMA. PRMT10 and 11 have been identified as putative PRMT genes,15−17 but their methylation activity has not yet been shown. As the founding member of the mammalian PRMT family, PRMT1, which was the first type I PRMT that was cloned and characterized, is the predominant arginine methyltransferase in most mammalian cells.18 Although extensive investigations have attempted to understand the biological importance of PRMTs, the knowledge of Received: July 20, 2015

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communication pathways between the dimerization arm and the catalytic site and on the subtle conformational modifications induced by dimerization. To gain mechanistic insight into the conformational fluctuations and allosteric pathways in PRMT structures, we initiated simulation studies of human PRMT1, a ubiquitously expressed member of the PRMT family that has been estimated to contribute more than 90% of the type I PRMT activity observed in cells and tissues.18 In our study, we modeled the monomer and dimer systems of hPRMT1 in both the presence and absence of the cofactor SAM and/or the R3 substrate. The dynamic conformational changes induced by dimerization and SAM binding were captured through molecular dynamics (MD) simulations. In addition, cross-correlations between the N-terminal region and the dimerization arm were observed, suggesting the existence of communication pathways. This long-distance communication is achieved through what is classically known as an allosteric regulation mechanism, which was theoretically elucidated through network topological analysis. Subsequently, six residues involved in the theoretical communication pathways were selected for a subsequent mutational study. The mutagenesis data demonstrate that the PRMT1 binding affinity with the cofactor SAM decreased progressively with a reduction in PRMT1 dimer formation, accordingly resulting in reduced methylation activity. Taken together, the findings obtained in this study underline the importance of PRMT1 dimer formation in SAM binding and catalytic methylation and reveal the allosteric regulation induced upon dimerization, which is conserved in PRMTs, thereby providing valuable information for the design of allosteric effectors against PRMTs.

the fundamental biochemistry of these enzymes remains limited. To elucidate the detailed relationship between the structure and function of PRMTs, several questions should be addressed. First, in the resolved crystal structures of PRMTs, the N-terminal region is mostly disordered due to its flexibility, similarly to the findings observed in SAH-PRMT1 structures (PDB entries 1OR8, 1ORH, and 1ORI) 8 and apoCARM1140−480 structures (PDB entries 3B3G and 2V7E).19 However, in the crystal structures of SAH-PRMT3 (PDB entry 1F3L)9 and SAH-CARM1140−480 (PDB entries 3B3F and 2V74),19 the N-terminal region is structured into three helices, namely αX, αY, and αZ, and helices αX and αY act as a lid that shields the cofactor SAH from the solvent. Surprisingly, in the apo-CARM128−507 structure (PDB entry 3B3J),19 part of helix αX has been transformed into strand β0, and the kink between helices αY and αZ has disappeared, yielding an inactive conformation. This finding suggests the existence of a molecular switch between the active and inactive conformations in the N-terminal region, specifically due to the presence of the above-mentioned kink between helices αY and αZ. However, the localization of a few molecules of benzamidine in this inactive structure may drive structural modifications in the N terminus to yield an unnatural conformation. Although the deletion of helix αX in rat PRMT1 has demonstrated its important roles in both SAM binding and catalysis,8 the conformational changes that occur in the N-terminal region during methylation catalysis should be further resolved. Moreover, the correlation of the N-terminal region with catalytic activity remains to be elucidated. Second, the dimerization arm, which is not well conserved in the PRMT family, is important in modifying the relative orientations of two monomers in dimerization. Although the mutant in which the entire dimerization arm has been deleted does not exhibit binding affinity with SAM and presents a complete lack of enzymatic activity,8,20 it is difficult to imagine its original structure remained, thus calling into new question the importance of the dimerization arm in dimer formation and SAM binding. In addition, dimer formation is a conserved feature among the PRMT family, and no crystal structure in the monomeric state has been resolved to date. However, little biochemical information is available to explain the relationship between dimer formation and methylation activity. If dimer or oligomer formation is a prerequisite for PRMT methylase activity, allosteric communication pathways that transmit any structural changes in the dimer interface to the active site could exist to facilitate methylation catalysis. The hypothesis that there is an allosteric pathway involving physically interconnected or thermodynamically linked residues through which signals are communicated has been supported by many studies.21 Allosteric regulation could occur in the absence of large conformational changes, and in this process, a protein would have the ability to facilitate allosteric pathways attributed to the small vibrations inherent to the protein structure and not due to a change in shape or fold.22 The protein structure network (PSN) is a powerful tool for probing allosteric function and has been successfully employed to probe the allosteric pathways in tRNA:protein complexes,23 methionyltRNA synthetase,24 tryptophanyl-tRNA synthetase,25 M2 muscarinic receptors,26 and Dronpa and a DNA clamp.27 In the resolved crystal structures of PRMTs, the dimer interface is formed between the dimerization arm and the outer surface of the SAM-binding site, including the N-terminal region. Hence, we focused on the characterization of the allosteric

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METHODS Homology Modeling. The sequence identity between hPRMT1 and rat PRMT1 is 99.7%, with the difference in only one residue (His161 in rat corresponds to Tyrosine in human). Hence, we modeled the hPRMT1 structure from the resolved rat PRMT1 structure (PDB entry 1OR8)8 by mutating His161 to tyrosine. For the unresolved helix αX, the CARM1 structure (PDB entry 3B3F)19 has high N-terminal-sequence similarity with PRMT1 (Figure S1A), especially for the signature sequence referred to as motif I (Y35F36xxY39) (Supporting Information). Hence, helix αX in CARM1 was used as the template, and homology modeling was performed using the MODELER program in the Discovery Studio 3.0 package (Accelrys Inc., San Diego, CA, USA). The optimized PRMT1 model was subjected to the stereochemical evaluation by PROCHECK. After verifying the rationality (Figure S1E), models of PRMT1D, symmetric PRMT1D‑SAM, and symmetric PRMT1D‑SAM‑R3 were subsequently constructed from PRMT1M according to the chain orientations and the position of SAM in 1OR8. The R3 peptide was also constructed based on the Cα atoms in the crystal structure to ensure that the target arginine was well situated in the pocket during the simulations. The side chains were then rotated manually to eliminate any visible spatial clashes. The protonation and ionization states of the models at pH 6.5−7.5 were calculated using the H++ system.28 The antechamber program was used to assign atomic charges and atom types to the cofactor SAM in GAFF (the general AMBER force field).29 All four models were obtained through four-step restrained energy minimizations to refine the stereochemistry using the steepest descent method in a vacuum with AMBER 12 software.30 B

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Journal of Chemical Information and Modeling Molecular Dynamics Simulations. The molecular dynamics (MD) simulations were performed using the GROMACS 4.3 package31 under an AMBER99SB force field32 with the explicit TIP3P water model. The parameters for SAM were described in our previous publication.33 A cubic box located within 10 Å of the protein was generated to bathe the molecule in a 0.15 mol/L NaCl solution. During the simulation, periodic boundary conditions were used in all of the calculations to avoid edge effects. The particle-mesh Ewald method was adopted to calculate the long-range electrostatics, and a cutoff radius of 10 Å was applied to any Leonard-Jones interaction. A velocity rescaling scheme was used to generate a constant-temperature system at 300 K, and Berendsen pressure coupling was used to equilibrate the systems during the simulation. The normalized covariance (correlation) analysis and standard PCA of the MD simulations were subsequently performed using CARMA software.34 The free-energy landscape of the conformational ensemble produced by MD simulations was represented in a two-dimensional (2D) representation, in which principal components PC1 and PC2 were selected as the reaction coordinates. The energy landscape along the two reaction coordinates was calculated by the function in eq 1, G(q1 , q2) = −kBT ln P(q1 , q2)

The length of path Dij between distant nodes i and j is the sum of the edge weights between the consecutive nodes (k, l) along the path, Dij =

k ,l

(eq 1)

(eq 2)

The correlation value Cij was derived from fluctuations information (eq 3), Cij =

Δγi (⃗ t ) ·Δγj(⃗ t ) ( Δγi (⃗ t )2

Δγj(⃗ t )2 )1/2

(eq 4)

The shortest distance Dij between all pairs of nodes was found using the Floyd−Warshall algorithm. Protein Expression and Purification. rPRMT1 (residues 11−353) and nine other mutants with an N-terminal 6X-His tag (MGHHHHHH), which was added to facilitate protein purification, were expressed in Escherichia coli BL21 (DE3) cells and purified using a HisTrap HP column followed by a desalting column (GE Healthcare). Peptide Synthesis. The sequence of the biotinylated NH2terminal 20-aa peptide of histone H4, denoted H4(1−20) _BTN, is Ac-SGRGKGGKGLGKGGAKRHRK(Biotin), and in this peptide, biotin is connected to the side-chain amino group of Lys20. The peptides were synthesized using standard solidphase peptide synthesis protocols, purified by C-18 RP-HPLC, and confirmed through MALDI-MS as described previously.39−42 Enzymatic Activity Assay. To evaluate the enzymatic activity of each mutant, a scintillation proximity assay (SPA)43 was performed in a 96-well plate at room temperature (approximately 25 °C). The reaction buffer contained 50 mM HEPES (pH 8.0), 1 mM EDTA, 50 mM NaCl, and 0.5 mM dithiothreitol (DTT). To analyze the tested samples, a 24-μL mixture of [3H]-labeled SAM (PerkinElmer, 18 Ci/mmol) and H4(1−20)_BTN peptide was incubated at room temperature for 5 min, and 6 μL of enzyme was then added to initiate the reaction. The final concentrations of SAM, the peptide, and the enzyme in the reaction mixture were 0.5 μM, 1 μM, and 20 nM, respectively. After 8 min, the reaction was quenched with 30 μL of isopropanol and then mixed with 10 μL of streptavidincoated SPA beads (PerkinElmer, dissolved in ddH2O, 20 mg/ mL). The products were analyzed using a Microbeta2 scintillation counter. For the negative control, 6 μL of reaction buffer was used instead of the enzyme. The activity of wild-type PRMT1 was normalized unity. All the data were averaged from duplicate results. Stopped-Flow Fluorescence Assay. The transient kinetics of the binding between SAM and the enzymes were studied through stopped-flow spectrometry (SX20, Applied Photophysics). One drive syringe contained 0.8 μM enzyme, and the other syringe contained various concentrations of SAM (0−128 μM, Sigma-Aldrich). Both of these species were dissolved in reaction buffer. After equal volumes of the two components were mixed in a 20-μL observation cell at 20−21 °C, the enzyme Trp fluorescence change obtained upon the binding of SAM to the enzyme was determined using an excitation wavelength of 295 nm and a wavelength cutoff of ≥320 nm. The widths of the entrance and exit slits of the monochromator were set to 2 mm, corresponding to a 9.3 nm bandpass. Averaged data from four to six shots were fitted with a single exponential function (eq 5), where a refers to the amplitude of the signal change and b is the observed rate kobs. The variable kobs was then plotted as a function of the SAM concentration and fitted with eq 6, where kon and koff are the association and dissociation rate constants of the SAM−enzyme binding reaction, respectively.

in which kB is the Boltzmann constant, T is the temperature of simulation, and P(q1, q2) is the normalized joint probability distribution. The energy surfaces from the raw data were smoothed by the kernel density smooth method encoded in the R program. Graphic views of the energy surface were generated by the RGL module in the R program as well. The detailed method was described in our previous publications.35,36 Communication Pathway Analysis. Communication pathways in PRMT1 dimerization were proposed through the combination of two modes based on the network topology of the protein. First, the protein structure network (PSN) is defined as a set of nodes connected by edges, in which each node represents an amino acid residue and each edge corresponds to a relationship between two residues. The PSNs were constructed using the RING server based on the average structures obtained from the MD simulations. In these networks, the edges represent physicochemical interactions weighted by Van der Waals contact score. The shortest pathway between the correlated residues was calculated using RINalyzer37 and the ShortestPath plugin of Cytoscape.38 Second, a dynamic network model, which is considered as weighted PSN, was set up using the VMD software. In this weighted network, the edges connect nodes only if the heavy atoms from two residues are within a distance of 4.5 Å in at least 75% of the snapshots analyzed,23 and the weight (Wij) of an edge between nodes i and j indicates the probability of information transfer, which was calculated based on the correlation values using the following equation (eq 2), Wij = −log(|Cij|)

∑ Wkl

F = a exp( −bt ) + c

(eq 3) C

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Figure 1. (A) Root mean square fluctuations (RMSFs) of Cα obtained from 200 ns MD simulations of the PRMT1M, PRMT1D, PRMT1D‑SAM, and PRMT1D‑SAM‑R3 models. For comparison, the RMSF value calculated from experimentally derived B factors of the rat PRMT1 crystal structure is also displayed. Helices αY-αZ and αA-αB and the dimerization arm are labeled with green, orange, and pink boxes, corresponding to the color coded in the structure presentation. All the charts were made with the R program. (B) H-bond and hydrophobic interactions involved in the dimer interface. The N-terminal region is colored green, the Rossmann fold is colored orange, the β barrel is colored blue, and the dimerization arm is colored pink. All the structural representations were made with Pymol1.3 software. (C) Left, energy landscape of the conformational ensemble of PRMT1M. The reaction coordinates were defined according to the PC1 and PC2 obtained from the PCA. Right, typical snapshots of PRMT1M extracted from the MD simulations display the flexible dimerization arm.

kobs = kon[SAM] + koff

The Dimerization Arm in the PRMT1M Displays Great Flexibility. Comparisons of the PRMT1 models helped identify the key elements underlying the dynamic dimerization and SAM binding of PRMT1. Based on the RMSD values monitored throughout the MD simulations (Figure S2A), the PRMT1M model possessed more flexible characteristics than the dimer models. The monitored RMSF values (Figure 1A), which reflect the consistency of the crystal structure with the dimer models, indicated the increased ability of PRMT1 to form a dimer in solution. In PRMT1M, the dimerization arm displayed great flexibility, and this flexibility was identified as the main factor contributing to the instability of the protein observed in the MD simulations. In addition, the solventexposed faces of the Rossmann-fold helices (αY, αZ, αA, and αB) also displayed more flexibility in the PRMT1M model compared with the dimer models. Hence, the dimer interface comprising the dimerization arm of one monomer and the four helices (αY, αZ, αA, and αB) of the other monomer could be stabilized in the dimer models. Because the dimerization arm in PRMT1M exhibits great conformational fluctuations, PCA was performed to investigate the collective motions of PRMT1M. The first two principle component PCs, which accounted for 45.7% and 45% of the overall motions, respectively, consisted of notable motions of the residues involved in the dimerization interface (Figure S3). Especially for the dimerization arm, it tended to bend toward the β-barrel domain and caused the whole structure to stabilize at a more compressed conformation (Movie S1), yielding an inactive conformation of PRMT1 in the monomer state. Based

(eq 6)

Simulation of the Data from the Stopped-Flow Experiment. The global simulation of the transient kinetics data was performed using KinTek Explorer (version 4.0, KinTek Corporation). The data sets for each enzyme that were obtained with different concentrations of SAM were fitted simultaneously to the predefined single-step binding model. The initial estimates of each parameter (e.g., initial intensity, amplitude, rates, and concentrations) were set according to the original data. Native PAGE. Native PAGE was used to determine the native protein oligomeric states. The mutants were prepared in a nondenaturing sample buffer (30 mM Tris-HCl, 10% glycerol, and 0.5% bromophenol blue), which maintained the secondary structure and native charge density of the proteins. The mutant proteins were loaded on the gel at 2 μg per lane and stained according to a standard Coomassie-blue protocol.

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RESULTS The catalytic core domain is well conserved in sequence (and therefore structure) among all PRMT members (Figure S1A). In this study, we modeled the hPRMT1 dimer with the cofactor SAM and the R3 substrate (Figure S1B) based on the resolved structure of rat PRMT1. To investigate the dynamics of dimer formation, we modeled the following four systems using MD simulations: the PRMT1 monomer (PRMT1M for short), the PRMT1 dimer (PRMT1D), the PRMT1 dimer in complex with SAM (PRMT1D‑SAM), and the PRMT1 dimer in complex with SAM and R3 (PRMT1D‑SAM‑R3). D

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Figure 2. (A) Secondary structures for the PRMT1M, PRMT1D, and PRMT1D‑SAM models as a function of time. The profile of the secondary structure transformation in the trajectories was calculated using DSSP. (B) Angle between helix αY and helix αZ in the PRMT1M and PRMT1D models as a function of the simulation time. (C) Detailed H-bond interactions in the PRMT1M, PRMT1D, and PRMT1D‑SAM models.

The Dynamics of the N-Terminal Region Facilitate SAM Binding upon Dimerization. Because of the structural conformational changes observed in the PRMT structures, a secondary structure prediction analysis was performed specifically for the N-terminal region (Figure 2A). Consistent with the resolved crystal structures, the N-terminus displayed extraordinary flexibility and became partially disordered in MD simulations. However, the N-terminus in PRMT1D‑SAM was relatively stable compared with the PRMT1D model. Since the N-terminus was not resolved in the rat PRMT1-SAH structure, the observation here may be attributed to the different effects caused by the cofactor SAM or product SAH (with methylated arginine substrate). On the other hand, the insufficient sampling in MD simulations might be another reason for this observation. Considering the participation of the N-terminus in SAM binding sites, its extreme flexibility is considered to play potential roles in methylation catalysis. In rat PRMT1, deleting helix αX reduced the SAM binding of the protein and abolished its enzymatic activity, suggesting the important roles of this terminus in both SAM binding and catalysis.8 Besides, the interactions between SAM (or R3 substrate) and the Nterminus are assumed to have potential influence on its conformation and dimer formation. This conclusion was supported by the previous finding that SAM binding can enhance dimerization by decreasing the Kd value of the monomer-to-dimer conversion.44 In the Rossmann-fold domain, the SAM-binding site is covered by the N-terminal helices αX and αY, which act as a lid that regulates the entrance of SAM. Helix αZ, located on the top of the active site, is also involved in the SAM-binding interactions. Because the kink between helices αY and αZ was absent in the apo-CARM128−507 structure (PDB entry 3B3J),19 which represents an unnatural inactive conformation, we monitored the angle between helices αY and αZ in PRMT1M

on the conformations from MD simulation and PCA analysis, a rough energy landscape (Figure 1C) for the conformational transition of PRMT1M projected onto the first two PCs was constructed. In the energy landscape, two typical deep wells, which reflect two low-energy-state conformations, were identified: one represents the PRMT1M conformation in which the dimerization arm became closer to the β-barrel domain, and the other represents the further compressed conformation of PRMT1M with the curved arm. In MD simulations, the dimerization arm was unable to remain in the extended state when exposed to the solvent in monomeric form, which was attributed to its highly hydrophobic outer surface. In the dimer models, multiple van der Waals interactions occurred in the dimer interface between a subunit (Leu49, Thr55, Phe63, Thr81, Ile83, Met86, Phe87, and Ile111) and its partner (Trp197, Trp198, Val201, Tyr202, Phe204, Met206, Cys208, and Ile209) (Figure 1B), which are mostly conserved in the resolved crystal structures.8,9 In addition to multiple hydrophobic interactions, the H-bonds obtained in the dimer interface were monitored throughout the MD simulations (Table S1). Of note, residues His196 and Asp205 in the dimerization arm form stable H-bonds with Glu46′ in helix αY and Asn115′ in the Rossmann fold (Figure 1B) with occupancies of 99.4% and 99.2%, respectively. These hydrophobic and H-bond interactions enhance the contacts between the dimerization arm and the Rossmann fold and thus become the main contributor to the structural stability of the dimer models. Experimental observations have shown that the destruction of the H-bond between the conserved residues Asn230 and Asp323 in the CARM1 S229E mutant (corresponding to Asn115 and Asp205 in PRMT1) impacts the dimerization, likely also resulting in diminished SAM binding and enzymatic activity.10 E

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Figure 3. (A) Dynamic cross-correlation map (DCCM) representing the collective atom fluctuations for the PRMT1D model. The strong (Cij = ±(0.5−1.0)) and weak (Cij = ±(0.3−0.5)) correlations are presented in red and gray, respectively. For clarity, the upper and lower triangles correspond to positive and negative correlations, respectively. Residues 28−50 correspond to helixes αX and αY (αXY), residues 51−174 correspond to the conserved Rossmann fold, residues 191−213 correspond to the dimerization arm (DA), and the remaining residues correspond to the β-barrel domain. (B) Representation of the allosteric communication pathways between the dimerization arm and the N-terminal region in PRMT1 in a single monomer (C) and in the dimer (D). The residues participating in the communication pathways are shown as sticks.

and PRMT1D during 1000 ns MD simulations (Figure 2B). Interestingly, in PRMT1D, the angle between helices αY and αZ was markedly enlarged compared with that found in PRMT1M. The enlarged angle likely corresponds to an enlarged SAMbinding site (Figure S2B), indicating that PRMT1D likely represents an intermediate in the dimerization process to facilitate subsequent SAM binding. To probe the molecular basis for the N-terminal fluctuations, an H-bond interaction analysis was subsequently conducted (Table S2). We discovered that an H-bond interaction network was stably formed in helix αX of PRMT1M (Figure 2C, left). Most importantly, residue Gly43, which is located between helices αX and αY, forms an H-bond with Ser103, and His45, which is located in helix αY, forms an H-bond with Glu100. Both of these H-bonds, acting as an anchor between the Nterminal region and the Rossmann fold, maintain helix αX in a straight conformation and the SAM-binding site in a closed state. However, in PRMT1D, almost no stable H-bonds were formed in the N-terminal sequence of one monomer. Instead, H-bond interactions were generated in the dimer surface, mostly between helices αY and αZ of one monomer and the dimerization arm and C-terminal region of the other monomer (Figure 2C, middle). For example, H-bonds were formed between Glu46 and His196′, between Glu46 and Arg351′, between Gly43 and Glu216′, between His41 and Asp349′, and between Asp37 and Lys324′. This finding, combined with steric clash, results in the N-terminal region of PRMT1D displaying less stability. Hence, the conformational fluctuations in the N terminus of PRMT1D induced by dimerization make the active binding site more accessible to SAM for binding, whereas in PRMT1D‑SAM and PRMT1D‑SAM‑R3, a series of H-bond and

hydrophobic interactions between cofactor SAM (Figure 2C, right) and the N-terminal region were formed, and the detailed analysis is described in the Supporting Information. The Correlation Analysis Results Indicate the Existence of Allosteric Communication Pathways. To explore the molecular basis for the dynamic interconnection of the dimerization arm and the N-terminal region, the crosscorrelation matrix of atomic fluctuations was measured over the course of the MD simulations. In PRMT1D, in addition to the local correlations along the diagonal line, correlations between distant parts were obtained and are labeled in boxed regions in Figure 3A. The motion of the Rossmann fold in one monomer was found to be anticorrelated to the β barrel in the other monomer, and this finding is likely attributable to the formation of the dimer interface. Most significantly, the Nterminal region is dynamically correlated to (with correlation | Cij| values ≥ 0.5) the motion of the dimerization arm in the same and the other monomers, even though these regions are located 34 Å/20 Å apart, as shown in the red box (Figure 3A and 3B). The degree of these similar correlation trends was significantly strengthened in the PRMT1D‑SAM‑R3 model (Figure S4), probably attributed to the binding with SAM and R3 substrate. These correlations between distant regions provide evidence of the presence of allosteric communication pathways in the PRMT1 dimerization process. As mentioned above, PRMT1D is considered the intermediate state for SAM binding and subsequent catalytic methylation. In this study, we selected residues located in the N-terminal region and the dimerization arm as the end points and identified communication pathways in PRMT1D through network topology, as listed in Table S4. Based on previous mutagenesis studies and structural F

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Journal of Chemical Information and Modeling significance, two theoretical communication pathways are proposed. In one monomer, the typical pathway transmits signals through residues His45-Met48-Glu144-Trp294-Val219Arg353(Arg351)-Gln189-Lys191-Lys194 (Figure 3C). In contrast, between the dimer interface, the communication pathway involves residues His41-Gly43-His45-Ile104-Tyr107/Ala108Ile111-Asn115-Asp205′ (Figure 3D), which incorporates the stable H-bond between Asn115 and Asp205′ observed in the MD simulations. Upon dimerization, the dimerization arm is accordingly stabilized, and these pathways are hypothesized to transmit the structural modifications that occur in the dimer interface to the SAM-binding site, facilitating subsequent methylation catalysis. The pathway analysis revealed the stable existence of a conserved subcommunication pathway involving Met48Glu144/Arg54-Trp294-Val219-Arg353(Arg351), indicating its notable effect in long-range communication. In this pathway, the mutation of Met48 (M48L and M48A) results in a loss of methylation activity and a change in the mono- and dimethylated products.45,46 The carboxylate group of Glu144 may act as the proton acceptor in arginine deprotonation,33 and a previous mutation experiment revealed the critical role of this group in methylation catalysis.8 In this study, Trp294, which is located in the highly conserved THW loop near the substratebinding groove, was hypothesized to also play a crucial role in the signal transduction to facilitate catalytic methylation. In addition, the negatively charged C-terminal carboxyl group of Arg353 (or Arg351) may play an important role in the binding of positively charged substrates,8 which also underscores the potential value of this allosteric pathway. To further explore the key residues involved in the communication pathways, mutation experiments were subsequently conducted. Mutation of Residues in the Allosteric Pathways Impairs PRMT1 Dimer Formation, SAM Binding, and Enzymatic Activity. The residues involved in key structural interactions in the pathways were mutated to alanine, and the resulting methylation activity was tested through in vitro enzymatic assays. Consistent with our prediction, the mutants displayed markedly reduced methylation activity compared with the wild-type enzyme (Figure 4A). Notably, three double mutants (H41A/D205A, H45A/W294A, and H45A/R353A), which were hypothesized to exert marked effects on the theoretical communication pathways, presented greatly diminished methylation activity. To further analyze the mechanism underlying the activities of the mutants, analyses of the dimerization status and SAM binding were subsequently performed through native PAGE and stopped-flow fluorescence assays, respectively. In the native PAGE, dimer formation was markedly reduced in the H45A, D205A, H41A/D205A, H45A/K194A, and H45A/R353A mutants (Figure 4B). Dimerization could barely be observed between these mutants, and the other mutants showed decreases to different degrees compared with the wild-type. In the stopped-flow assay, the time-dependent Trp fluorescence change was observed upon SAM binding and was harnessed to determine the transient kinetics. A singleexponential function was adequate to fit the data trace (data points evenly distributed around the fitting curve). The plot of the obtained kobs as a function of the SAM concentration revealed a linear relationship, indicating a one-step binding event (Figure S5A, exemplified by rPRMT1(H41A)). The kon (association rate constant) and koff (dissociation rate constant) values were estimated from the slope and the y-intercept of the

Figure 4. (A) Enzymatic activity of each mutant determined through radiometric assays. (B) Native-PAGE of each mutant.

regression line. However, the value of koff calculated through this conventional analysis is considered inaccurate. Errors may also arise from fluorescence fluctuations caused by the equipment used. Therefore, more accurate kon and koff values (Table 1) were obtained through a global simulation, which was performed using KinTek ExplorerTM46 with a predefined one-step model and pregained estimates of the parameters (Figure S5B). Table 1. Measurement of kon, koff, and Kd Values through a Global Simulation of the Data from the Stopped-Flow Assay kon (μM‑1 s‑1) wt H41A H45A K194A D205A W294A H41A/D205A H45A/R353A H45A/W294A a

0.20 0.30 -a 0.07 0.08 0.18 -

± 0.006 ± 0.008 ± 0.005 ± 0.005 ± 0.020

koff (s‑1) 2.75 4.38 2.50 1.45 8.17 -

± 0.091 ± 0.139 ± 0.143 ± 0.098 ± 0.929

Kd (μM)b 13.8 ± 0.87 14.6 ± 0.85 35.7 ± 4.59 18.1 ± 2.36 45 ± 10.2 -

No signal change. bCalculated as koff/kon.

The Kd (13.8 μM) of SAM for wild type PRMT1 was in agreement with the Kd (corresponding to Ki(SAM) in the paper) calculated from steady-state kinetic experiment.47 The analysis of single mutants revealed that no signal changes were obtained with H45A and D205A, indicating a lack of binding events with the SAM concentration used in the assay. Correspondingly, both mutants remained mostly in the monomeric state (Figure 4B, lanes 2 and 4), as determined by native PAGE, indicating G

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Figure 5. Proposed models for flexible PRMT1M, intermediate PRMT1D, and catalytic PRMT1D‑SAM. These models emphasize the allosteric communication pathways between the N-terminal region and the dimerization arm in PRMT1 dimerization, which facilitates SAM binding and subsequent catalytic methylation.

the structural fold of PRMTs makes graph theory a powerful tool for probing these allosteric pathways. In the end, the preparation of mutants with decreased methylation activity validates the essential roles of the residues involved in the communication pathways. These findings in combination with the results from native PAGE and SAM-binding experiments demonstrate a qualitative correlation between dimer formation and SAM binding. We conclude that the dimerization of PRMT1 facilitates SAM binding and thus is essential for methylation activity. Although most mutations on the dimer interface could directly influence the enzymatic activity, the mutation (W294A), apart from the dimer interface, also displayed reduced enzymatic activity. This observations support the notion of long-distance regulation, which mediates the correlation between the active site and the dimerization interface. In addition, the residues involved in cofactor SAM binding and the intermolecular contacts in the putative dimer interface and along the communication pathways are evolutionarily conserved. As a result, we speculate that the allosteric communication proposed in PRMT1 may also be found in the entire PRMT family. In epigenetic regulation, protein arginine (PRMT) and protein lysine (PKMT) methyltransferases catalyze the transfer of methyl groups from SAM to arginine and lysine on substrates, respectively. The discovery of small-molecular inhibitors targeting PRMTs and PKMTs is of great value in the investigation of the biological function and therapeutic potential of these enzymes.6,49−56 Hence, the selectivity of these inhibitors becomes the real challenge. In this study, the allosteric mechanism proposed may be an effective alternative for selective drug design because it presents specific advantages compared with strategies targeting cofactor SAM- or substratebinding pockets. In PRMT3, inhibitors that bind at the allosteric site located at the interface between the two subunits of the homodimer have been identified, confirming the feasibility of mechanism of action (MOA)-based drug discovery.57,58 Compared with conventional orthosteric drugs for specific protein targets, allosteric drugs have fewer or reduced side effects due to their improved selectivity.59 Considering the allosteric site, residues Arg396, Glu422, and Thr466 in PRMT3 correspond to residues Lys194, Asp220, and His264 in PRMT1, respectively. Of these, residue Lys194 is theoretically involved in the allosteric communication pathway of PRMT1, hinting that allosteric regulation is reasonable and that an allosteric pocket also exists in PRMT1. Although the allosteric communications and inhibitors have been investigated in PRMTs family,19,57,58 the detailed allosteric communication

that dimerization is indeed important for SAM binding. As mentioned above, the side chain of His45 forms H-bonds with the ribose of cofactor SAM, and Asp205 is involved in stable Hbond interactions with the Asn115′ residue in the other monomer. The decreases in dimer formation and SAM-binding affinity may be attributed to the destruction of these interactions in the allosteric regulation. Lys194, located in the dimerization arm, forms a H-bond with Glu46′ in the other monomer. The K194A mutant also exhibited reduced dimer formation (Figure 4B, lane 3) and SAM-binding affinity (Kd = 35.7 μM). The H41A and W294A mutations had little effect on SAM binding (with Kd values of 14.6 μM and 18.1 μM, respectively), and dimer formation was almost unchanged (Figure 4B, lanes 1 and 5). Accordingly, the K194A, H41A, and W294A mutants displayed limited decreases in enzymatic activity compared with the wild-type enzyme. The Kd values resulting from the stopped-flow assay were consistent with the dimerization status obtained through native PAGE (lower Kd value corresponds to increased dimer formation), indicating that dimerization is indeed important for SAM binding. Our observations support the hypothesis that dimer formation is essential for cofactor SAM binding and catalytic methylation, which corresponds to our theoretical prediction and has just been proved in yeast Hmt1 research.48

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DISCUSSION The findings obtained in this study support the regulatory roles of the dimerization arm and the N-terminal region in methylation activity and provide a structural basis for the functional characterization of the PRMT family. Three structural features characterize the dynamic progression in PRMT catalysis. First, the dimerization arm in PRMT1M displays marked conformational fluctuation when exposed to solvent, whereas the N-terminal region is relatively stable with the active site in an enclosed state, theoretically indicating that the monomer is found in an unnatural inactive state (Figure 5, left). Second, once the arm is stabilized by dimer formation, the N-terminal region becomes disordered and flexible, with the active site in an enlarged state, indicating that PRMT1D is a reasonable intermediate for cofactor SAM binding (Figure 5, middle). Third, based on the dynamic correlation between the dimerization arm and the N-terminal region, allosteric communication pathways were elucidated theoretically, which facilitates SAM binding and subsequent catalytic methylation (Figure 5, right). Although no significant coupling motion between the N-terminal region and the dimerization arm was captured in the MD simulations, their inherent correlation in H

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Journal of Chemical Information and Modeling pathways proposed here for the first time are basically theoretical results, which need more sophisticated experiments for further confirmation. Besides, the relationship of R3 substrate binding with dimerization and SAM binding also needs further research. Nevertheless, the allosteric communication described is likely to be applicable to the entire PRMT family, which will enable further investigations of the physiological function of this family and shed new light on the rational design of inhibitors with both efficiency and specificity.

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(7) Cha, B.; Jho, E. H. Protein Arginine Methyltransferases (Prmts) as Therapeutic Targets. Expert Opin. Ther. Targets 2012, 16, 651−664. (8) Zhang, X.; Cheng, X. Structure of the Predominant Protein Arginine Methyltransferase Prmt1 and Analysis of Its Binding to Substrate Peptides. Structure 2003, 11, 509−520. (9) Zhang, X.; Zhou, L.; Cheng, X. Crystal Structure of the Conserved Core of Protein Arginine Methyltransferase Prmt3. EMBO J. 2000, 19, 3509−3519. (10) Higashimoto, K.; Kuhn, P.; Desai, D.; Cheng, X.; Xu, W. Phosphorylation-Mediated Inactivation of Coactivator-Associated Arginine Methyltransferase 1. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 12318−12323. (11) Kang, N.; Chen, P.; Chen, Y.; Zeng, H.; He, X.; Zhu, Y. Prmt6Mediates Cse Induced Inflammation and Apoptosis. Int. Immunopharmacol. 2015, 24, 95−101. (12) Dillon, M. B.; Rust, H. L.; Thompson, P. R.; Mowen, K. A. Automethylation of Protein Arginine Methyltransferase 8 (Prmt8) Regulates Activity by Impeding S-Adenosylmethionine Sensitivity. J. Biol. Chem. 2013, 288, 27872−27880. (13) Sun, L.; Wang, M.; Lv, Z.; Yang, N.; Liu, Y.; Bao, S.; Gong, W.; Xu, R. Structural Insights into Protein Arginine Symmetric Dimethylation by Prmt5. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 20538−20543. (14) Yang, Y.; Hadjikyriacou, A.; Xia, Z.; Gayatri, S.; Kim, D.; ZuritaLopez, C.; Kelly, R.; Guo, A.; Li, W.; Clarke, S. G.; Bedford, M. T. Prmt9 Is a Type Ii Methyltransferase That Methylates the Splicing Factor Sap145. Nat. Commun. 2015, 6, 6428. (15) Krause, C. D.; Yang, Z. H.; Kim, Y. S.; Lee, J. H.; Cook, J. R.; Pestka, S. Protein Arginine Methyltransferases: Evolution and Assessment of Their Pharmacological and Therapeutic Potential. Pharmacol. Ther. 2007, 113, 50−87. (16) Niu, L.; Lu, F.; Pei, Y.; Liu, C.; Cao, X. Regulation of Flowering Time by the Protein Arginine Methyltransferase Atprmt10. EMBO Rep. 2007, 8, 1190−1195. (17) Scebba, F.; De Bastiani, M.; Bernacchia, G.; Andreucci, A.; Galli, A.; Pitto, L. Prmt11: A New Arabidopsis Mbd7 Protein Partner with Arginine Methyltransferase Activity. Plant J. 2007, 52, 210−222. (18) Tang, J.; Frankel, A.; Cook, R. J.; Kim, S.; Paik, W. K.; Williams, K. R.; Clarke, S.; Herschman, H. R. Prmt1 Is the Predominant Type I Protein Arginine Methyltransferase in Mammalian Cells. J. Biol. Chem. 2000, 275, 7723−7730. (19) Troffer-Charlier, N.; Cura, V.; Hassenboehler, P.; Moras, D.; Cavarelli, J. Functional Insights from Structures of CoactivatorAssociated Arginine Methyltransferase 1 Domains. EMBO J. 2007, 26, 4391−4401. (20) Silver, P. A.; McBride, A. E.; Soriano, M. A.; Filman, D. J.; Weiss, V. H.; Hogle, J. M. The Structure and Oligomerization of the Yeast Arginine Methyltransferase, Hmt1. Nat. Struct. Biol. 2000, 7, 1165−1171. (21) Goodey, N. M.; Benkovic, S. J. Allosteric Regulation and Catalysis Emerge Via a Common Route. Nat. Chem. Biol. 2008, 4, 474−482. (22) Vanwart, A. T.; Eargle, J.; Luthey-Schulten, Z.; Amaro, R. E. Exploring Residue Component Contributions to Dynamical Network Models of Allostery. J. Chem. Theory Comput. 2012, 8, 2949−2961. (23) Sethi, A.; Eargle, J.; Black, A. A.; Luthey-Schulten, Z. Dynamical Networks in Trna:Protein Complexes. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 6620−6625. (24) Ghosh, A.; Vishveshwara, S. A Study of Communication Pathways in Methionyl- Trna Synthetase by Molecular Dynamics Simulations and Structure Network Analysis. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 15711−15716. (25) Hansia, P.; Ghosh, A.; Vishveshwara, S. Ligand Dependent Intra and Inter Subunit Communication in Human Tryptophanyl Trna Synthetase as Deduced from the Dynamics of Structure Networks. Mol. BioSyst. 2009, 5, 1860. (26) Miao, Y.; Nichols, S. E.; Gasper, P. M.; Metzger, V. T.; McCammon, J. A. Activation and Dynamic Network of the

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jcim.5b00454. Detailed analysis of the PRMT1 structure and interactions involved in the SAM-binding site (PDF) Movie showing the dynamical conformations of PRMT1 (MPG)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. *E-mail: [email protected]. Author Contributions ⊥

R.Z., Y.X., and H.H. contributed equally.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We are deeply indebted to the reviewers for the valuable comments that improved the results and discussion of MD simulations. This study was supported by the Ministry of Science and Technology of China (2015CB910304 to H.J.), the Hi-Tech Research and Development Program of China (2012AA020301 and 2012AA01A305 to K.Y. and 2012AA020302 to C.L.), the National Natural Science Foundation of China (81472378 to Y.H., 81430084 and 21472208 to C.L., 21203131 to G.H., and 81302700 to Z.L.), and NIH Grant (R01GM086717 to Y.G.Z.). The computation resources were supported by the Computer Network Information Center of the Chinese Academy of Sciences at Tianjin and the Shanghai Supercomputing Center.

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REFERENCES

(1) Bedford, M. T. Arginine Methylation at a Glance. J. Cell Sci. 2007, 120, 4243−4246. (2) Bedford, M. T.; Clarke, S. G. Protein Arginine Methylation in Mammals: Who, What, and Why. Mol. Cell 2009, 33, 1−13. (3) Wolf, S. S. The Protein Arginine Methyltransferase Family: An Update About Function, New Perspectives and the Physiological Role in Humans. Cell. Mol. Life Sci. 2009, 66, 2109−2121. (4) Yang, Y.; Bedford, M. T. Protein Arginine Methyltransferases and Cancer. Nat. Rev. Cancer 2013, 13, 37−50. (5) Spannhoff, A.; Sippl, W.; Jung, M. Cancer Treatment of the Future: Inhibitors of Histone Methyltransferases. Int. J. Biochem. Cell Biol. 2009, 41, 4−11. (6) Heinke, R.; Spannhoff, A.; Meier, R.; Trojer, P.; Bauer, I.; Jung, M.; Sippl, W. Virtual Screening and Biological Characterization of Novel Histone Arginine Methyltransferase Prmt1 Inhibitors. ChemMedChem 2009, 4, 69−77. I

DOI: 10.1021/acs.jcim.5b00454 J. Chem. Inf. Model. XXXX, XXX, XXX−XXX

Article

Journal of Chemical Information and Modeling M2Muscarinic Receptor. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 10982−10987. (27) Hu, G.; Yan, W.; Zhou, J.; Shen, B. Residue Interaction Network Analysis of Dronpa and a DNA Clamp. J. Theor. Biol. 2014, 348, 55− 64. (28) Gordon, J. C.; Myers, J. B.; Folta, T.; Shoja, V.; Heath, L. S.; Onufriev, A. H++: A Server for Estimating Pkas and Adding Missing Hydrogens to Macromolecules. Nucleic Acids Res. 2005, 33, W368− 371. (29) Wang, J.; Wolf, R. M.; Caldwell, J. W.; Kollman, P. A.; Case, D. A. Development and Testing of a General Amber Force Field. J. Comput. Chem. 2004, 25, 1157−1174. (30) Case, D. A.; Cheatham, T. E., 3rd; Darden, T.; Gohlke, H.; Luo, R.; Merz, K. M., Jr.; Onufriev, A.; Simmerling, C.; Wang, B.; Woods, R. J. The Amber Biomolecular Simulation Programs. J. Comput. Chem. 2005, 26, 1668−1688. (31) Hess, B.; Van Der Spoel, D.; Lindahl, E.; Kutzner, C. J. Chem. Theory Comput. 2008, 4, 435. (32) Lindorff-Larsen, K.; Piana, S.; Palmo, K.; Maragakis, P.; Klepeis, J. L.; Dror, R. O.; Shaw, D. E. Improved Side-Chain Torsion Potentials for the Amber Ff99sb Protein Force Field. Proteins: Struct., Funct., Genet. 2010, 78, 1950−1958. (33) Zhang, R.; Li, X.; Liang, Z.; Zhu, K.; Lu, J.; Kong, X.; Ouyang, S.; Li, L.; Zheng, Y. G.; Luo, C. Theoretical Insights into Catalytic Mechanism of Protein Arginine Methyltransferase 1. PLoS One 2013, 8, e72424. (34) Glykos, N. M. Software News and Updates. Carma: A Molecular Dynamics Analysis Program. J. Comput. Chem. 2006, 27, 1765−1768. (35) Ye, F.; Zhang, J.; Liu, H.; Hilgenfeld, R.; Zhang, R.; Kong, X.; Li, L.; Lu, J.; Zhang, X.; Li, D.; Jiang, H.; Yang, C. G.; Luo, C. Helix Unfolding/Refolding Characterizes the Functional Dynamics of Staphylococcus Aureus Clp Protease. J. Biol. Chem. 2013, 288, 17643−17653. (36) Du, Y.; Yang, H.; Xu, Y.; Cang, X.; Luo, C.; Mao, Y.; Wang, Y.; Qin, G.; Luo, X.; Jiang, H. Conformational Transition and Energy Landscape of Erbb4 Activated by Neuregulin1beta: One Microsecond Molecular Dynamics Simulations. J. Am. Chem. Soc. 2012, 134, 6720− 6731. (37) Doncheva, N. T.; Klein, K.; Domingues, F. S.; Albrecht, M. Analyzing and Visualizing Residue Networks of Protein Structures. Trends Biochem. Sci. 2011, 36, 179−182. (38) Smoot, M. E.; Ono, K.; Ruscheinski, J.; Wang, P. L.; Ideker, T. Cytoscape 2.8: New Features for Data Integration and Network Visualization. Bioinformatics 2011, 27, 431−432. (39) Xie, N.; Elangwe, E. N.; Asher, S.; Zheng, Y. G. A Dual-Mode Fluorescence Strategy for Screening Hat Modulators. Bioconjugate Chem. 2009, 20, 360−366. (40) Feng, Y.; Li, M.; Wang, B.; Zheng, Y. G. Discovery and Mechanistic Study of a Class of Protein Arginine Methylation Inhibitors. J. Med. Chem. 2010, 53, 6028−6039. (41) Feng, Y.; Xie, N.; Wu, J.; Yang, C.; Zheng, Y. G. Inhibitory Study of Protein Arginine Methyltransferase 1 Using a Fluorescent Approach. Biochem. Biophys. Res. Commun. 2009, 379, 567−572. (42) Hu, H.; Owens, E. A.; Su, H.; Yan, L.; Levitz, A.; Zhao, X.; Henary, M.; Zheng, Y. G. Exploration of Cyanine Compounds as Selective Inhibitors of Protein Arginine Methyltransferases: Synthesis and Biological Evaluation. J. Med. Chem. 2015, 58, 1228−1243. (43) Wu, J.; Xie, N.; Feng, Y.; Zheng, Y. G. Scintillation Proximity Assay of Arginine Methylation. J. Biomol. Screening 2012, 17, 237−244. (44) Lakowski, T. M.; Zurita-Lopez, C.; Clarke, S. G.; Frankel, A. Approaches to Measuring the Activities of Protein Arginine NMethyltransferases. Anal. Biochem. 2010, 397, 1−11. (45) Gui, S.; Wooderchak, W. L.; Daly, M. P.; Porter, P. J.; Johnson, S. J.; Hevel, J. M. Investigation of the Molecular Origins of ProteinArginine Methyltransferase I (Prmt1) Product Specificity Reveals a Role for Two Conserved Methionine Residues. J. Biol. Chem. 2011, 286, 29118−29126.

(46) Johnson, K. A.; Simpson, Z. B.; Blom, T. Global Kinetic Explorer: A New Computer Program for Dynamic Simulation and Fitting of Kinetic Data. Anal. Biochem. 2009, 387, 20−29. (47) Obianyo, O.; Osborne, T. C.; Thompson, P. R. Kinetic Mechanism of Protein Arginine Methyltransferase 1. Biochemistry 2008, 47, 10420−10427. (48) Li, H. T.; Gong, T.; Zhou, Z.; Liu, Y. T.; Cao, X.; He, Y.; Chen, C. D.; Zhou, J. Q. Yeast Hmt1 Catalyses Asymmetric Dimethylation of Histone H3 Arginine 2 in Vitro. Biochem. J. 2015, 467, 507−515. (49) Sinha, S. H.; Owens, E. A.; Feng, Y.; Yang, Y.; Xie, Y.; Tu, Y.; Henary, M.; Zheng, Y. G. Synthesis and Evaluation of Carbocyanine Dyes as Prmt Inhibitors and Imaging Agents. Eur. J. Med. Chem. 2012, 54, 647−659. (50) Dillon, M. B. C.; Bachovchin, D. A.; Brown, S. J.; Finn, M. G.; Rosen, H.; Cravatt, B. F.; Mowen, K. A. Novel Inhibitors for Prmt1 Discovered by High-Throughput Screening Using Activity-Based Fluorescence Polarization. ACS Chem. Biol. 2012, 7, 1198−1204. (51) Wang, J.; Chen, L.; Sinha, S. H.; Liang, Z.; Chai, H.; Muniyan, S.; Chou, Y. W.; Yang, C.; Yan, L.; Feng, Y.; Li, K. K.; Lin, M. F.; Jiang, H.; Zheng, Y. G.; Luo, C. Pharmacophore-Based Virtual Screening and Biological Evaluation of Small Molecule Inhibitors for Protein Arginine Methylation. J. Med. Chem. 2012, 55, 7978−7987. (52) Dowden, J.; Pike, R. A.; Parry, R. V.; Hong, W.; Muhsen, U. A.; Ward, S. G. Small Molecule Inhibitors That Discriminate between Protein Arginine N-Methyltransferases Prmt1 and Carm1. Org. Biomol. Chem. 2011, 9, 7814. (53) Selvi, B. R.; Batta, K.; Kishore, A. H.; Mantelingu, K.; Varier, R. A.; Balasubramanyam, K.; Pradhan, S. K.; Dasgupta, D.; Sriram, S.; Agrawal, S.; Kundu, T. K. Identification of a Novel Inhibitor of Coactivator-Associated Arginine Methyltransferase 1 (Carm1)-Mediated Methylation of Histone H3 Arg-17. J. Biol. Chem. 2010, 285, 7143−7152. (54) Feng, Y.; Li, M.; Wang, B.; Zheng, Y. G. Discovery and Mechanistic Study of a Class of Protein Arginine Methylation Inhibitors. J. Med. Chem. 2010, 53, 6028−6039. (55) Castellano, S.; Milite, C.; Ragno, R.; Simeoni, S.; Mai, A.; Limongelli, V.; Novellino, E.; Bauer, I.; Brosch, G.; Spannhoff, A.; Cheng, D.; Bedford, M. T.; Sbardella, G. Design, Synthesis and Biological Evaluation of Carboxy Analogues of Arginine Methyltransferase Inhibitor 1 (Ami-1). ChemMedChem 2010, 5, 398−414. (56) Spannhoff, A.; Machmur, R.; Heinke, R.; Trojer, P.; Bauer, I.; Brosch, G.; Schule, R.; Hanefeld, W.; Sippl, W.; Jung, M. A Novel Arginine Methyltransferase Inhibitor with Cellular Activity. Bioorg. Med. Chem. Lett. 2007, 17, 4150−4153. (57) Liu, F.; Li, F.; Ma, A.; Dobrovetsky, E.; Dong, A.; Gao, C.; Korboukh, I.; Liu, J.; Smil, D.; Brown, P. J.; Frye, S. V.; Arrowsmith, C. H.; Schapira, M.; Vedadi, M.; Jin, J. Exploiting an Allosteric Binding Site of Prmt3 Yields Potent and Selective Inhibitors. J. Med. Chem. 2013, 56, 2110−2124. (58) Siarheyeva, A.; Senisterra, G.; Allali-Hassani, A.; Dong, A.; Dobrovetsky, E.; Wasney; Gregory, A.; Chau, I.; Marcellus, R.; Hajian, T.; Liu, F.; Korboukh, I.; Smil, D.; Bolshan, Y.; Min, J.; Wu, H.; Zeng, H.; Loppnau, P.; Poda, G.; Griffin, C.; Aman, A.; Brown; Peter, J.; Jin, J.; Al-awar, R.; Arrowsmith; Cheryl, H.; Schapira, M.; Vedadi, M. An Allosteric Inhibitor of Protein Arginine Methyltransferase 3. Structure 2012, 20, 1425−1435. (59) Pei, J.; Yin, N.; Ma, X.; Lai, L. Systems Biology Brings New Dimensions for Structure-Based Drug Design. J. Am. Chem. Soc. 2014, 136, 11556−11565.

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