Pharmaceutical Sciences, Faculty of
DSpace
UBCV
Rowley, Michael James Anthony
Doctor of Philosophy - PhD
University of British Columbia
Arginine methylation is a prevalent post-translational modification in biology present on 0.5% of total arginine proteins in mammalian cells. Protein arginine methylation is a key epigenetic modification on histone proteins, that can promote or inhibit gene expression depending on the type and site of arginine methylation. Protein arginine N-methyltransferases (PRMTs) are the epigenetic writers responsible for arginine methylation and have emerged as therapeutic targets in several health disorders including neurodegenerative and cardiovascular disease, and cancer. PRMT2 was recently added to the list of epigenetic writers associated with oncogenic transcriptional programming, specifically in glioblastoma (GBM), hepatocellular carcinoma (HCC), and renal cell carcinoma (RCC). Unlike the other PRMT enzymes, the function of PRMT2 in cells remains enigmatic due to its lackluster in vitro methylation kinetics, making it difficult to characterize. The purpose of this research is to develop techniques for measuring PRMT2 interactions and to use these techniques to characterize how PRMT2 interacts with other proteins and histones. The development of the filter binding and phosphor screening (FBAPS) assay improved the throughput of the traditional filter binding assay used to characterize kinetic parameters of PRMT enzymes. Characterization of PRMT2 through filter binding assay revealed that exposing the radiolabeled S-adenosyl-L-methionine (SAM) cofactor to elevated temperatures for long periods of time create degradation products that confound analysis. We used differential scanning fluorimetry (DSF) to characterize PRMT2 thermal stability to identify conditions that may improve PRMT2 catalytic activity and to measure PRMT2 binding interactions. We identified several excipients that could thermally stabilize PRMT2 but did not improve catalytic activity. Additionally, we found that PRMT2 can interact with SAM and sinefungin (a SAM analogue) in a thermally stabilizing manner. Using kinetic and biophysical assays, we explored a noncatalytic role for PRMT2 in histone methylation by investigating interactions between PRMT2, histones, and other PRMTs. The result of this study indicated that PRMT2 interacts with PRMT1 in a cofactor- and domain-dependent manner. Further PRMT2 could modulate the histone methylation capacity of PRMT1 in a macromolecular-dependent manner. This work reveals the ability of PRMT2 to serve a noncatalytic role through its SH3 domain in generating site-specific histone methylation marks.
BIOCHEMICAL INVESTIGATIONS OF PROTEIN ARGININE N-METHYLTRANSFERASE 2 by Michael James Anthony Rowley B.Sc. (Hons), The University of Calgary, 2019 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pharmaceutical Sciences) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) May 2024 © Michael James Anthony Rowley, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: BIOCHEMICAL INVESTIGATIONS OF PROTEIN ARGININE N-METHYLTRANSFERASE 2 submitted by Michael James Anthony Rowley in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Pharmaceutical Sciences Examining Committee: Dr. Adam Frankel, Associate Professor, Pharmaceutical Sciences, UBC Supervisor Dr. Filip van Petegem, Professor, Biochemistry and Molecular Biology, UBC Supervisory Committee Member Dr. Martin Tanner, Professor, Chemistry, UBC University Examiner Dr. Judy Wong, Professor, Pharmaceutical Sciences, UBC University Examiner Dr. Muriel le Romancer, Professor, Centre de Recherche en Cancérologie de Lyon, Université de Lyon External Examiner Additional Supervisory Committee Members: Dr. Brent Page, Assistant Professor, Pharmaceutical Sciences, UBC Supervisory Committee Member Dr. Karla Williams, Associate Professor, Pharmaceutical Sciences, UBC Supervisory Committee Member iii Abstract Arginine methylation is a prevalent post-translational modification in biology present on 0.5% of total arginine proteins in mammalian cells. Protein arginine methylation is a key epigenetic modification on histone proteins, that can promote or inhibit gene expression depending on the type and site of arginine methylation. Protein arginine N-methyltransferases (PRMTs) are the epigenetic writers responsible for arginine methylation and have emerged as therapeutic targets in several health disorders including neurodegenerative and cardiovascular disease, and cancer. PRMT2 was recently added to the list of epigenetic writers associated with oncogenic transcriptional programming, specifically in glioblastoma (GBM), hepatocellular carcinoma (HCC), and renal cell carcinoma (RCC). Unlike the other PRMT enzymes, the function of PRMT2 in cells remains enigmatic due to its lackluster in vitro methylation kinetics, making it difficult to characterize. The purpose of this research is to develop techniques for measuring PRMT2 interactions and to use these techniques to characterize how PRMT2 interacts with other proteins and histones. The development of the filter binding and phosphor screening (FBAPS) assay improved the throughput of the traditional filter binding assay used to characterize kinetic parameters of PRMT enzymes. Characterization of PRMT2 through filter binding assay revealed that exposing the radiolabeled S-adenosyl-L-methionine (SAM) cofactor to elevated temperatures for long periods of time create degradation products that confound analysis. We used differential scanning fluorimetry (DSF) to characterize PRMT2 thermal stability to identify conditions that may improve PRMT2 catalytic activity and to measure PRMT2 binding interactions. We identified several excipients that could thermally stabilize PRMT2 but did not improve catalytic activity. Additionally, we found that PRMT2 can interact with SAM and sinefungin (a SAM analogue) in a thermally stabilizing manner. Using kinetic and biophysical assays, we explored a noncatalytic role for PRMT2 in histone methylation by investigating interactions between PRMT2, histones, and other PRMTs. The result of this study indicated that PRMT2 interacts with PRMT1 in a cofactor- and domain-dependent manner. Further PRMT2 could modulate the histone methylation capacity of PRMT1 in a macromolecular-dependent manner. This work reveals the iv ability of PRMT2 to serve a noncatalytic role through its SH3 domain in generating site-specific histone methylation marks. v Lay Summary Protein arginine N-methyltransferase 2 (PRMT2) is a member of the PRMT enzyme family that is involved in many cellular processes and has been implicated in the onset and severity of several health disorders, including neurodegenerative, cardiovascular, and neuromuscular disease, and cancer. While evidence suggests that PRMT2 is responsible for modifying specific DNA-associated proteins, its mechanism for how it works inside of cells and contributes to a disease phenotype has yet to be fully elucidated. This research identified new techniques that can be used to study PRMT2 and how it interacts with other proteins. Using these new techniques, we show that PRMT2 can directly interact with histones, the proteins around which DNA is wrapped, and other PRMT enzymes. Through these interactions, PRMT2 can change the activity of other PRMT enzymes towards histone proteins. These new findings indicate that PRMT2 might have a greater role in cells through these additional interactions with other PRMTs. vi Preface A version of Chapter 2 has been published and adapted here with permission from the publisher, Elsevier. â¢ Rowley, M. J., Taylor, S. A., Frankel, A. Protein arginine N-methyltransferase activity determination with filter binding and phosphor screening (FBAPS) assay. (2022) Anal. Biochem. 653. 1-7. I designed and conducted all experiments, parameterization, data analysis, and protein purification. Shona A. Taylor performed preliminary experiments, which does not appear in this thesis, and contributed helpful points of discussion. I wrote and edited the manuscript with Dr. Adam Frankel. The PRMT2-related content was not included in the published manuscript. Dr. Adam Frankel and I contributed to the experimental design. A version of Chapters 3 and 4 has been published and adapted here with permission from the publisher, Elsevier. â¢ Rowley, M. J., Prout-Holm, R. A., Liu, R. W., Hendrickson-Rebizant, T., Ige, O. O., Lakowski, T. M., Frankel, A. Protein arginine N-methyltransferase 2 plays a noncatalytic role in the histone methylation activity of PRMT1. (2023) J. Biol. Chem. 299(12). 1-18. I expressed and purified all recombinant proteins used in this study, performed all in vitro methylation assays and was responsible for the enzyme kinetics studies. Riley Austin Prout-Holm performed cloning on the PRMT2 SH3 domain into pET28a(+). Rui Wen Liu performed mutagenesis on PRMT2 to generate the PRMT2H112Q mutant. Thordur Hendrickson-Rebizant and Olufola O. Ige expressed and purified recombinant Xenopus laevis histone proteins used in preliminary experiments of this study. Dr. Ted M. Lakowski generated the GST-PRMT2ÎSH3, His6CARM1, His6-PRMT6 clones used in this study and contributed to editing the manuscript and experimental design. I wrote and edited the manuscript with Dr. Adam Frankel. Dr. Adam Frankel and I contributed to the experimental design. Please see footnotes at the beginning of each chapter for similar information. vii Table of Contents Abstract ....................................................................................................................................................... iii Lay Summary .............................................................................................................................................. v Preface ......................................................................................................................................................... vi Table of Contents ...................................................................................................................................... vii List of Tables ............................................................................................................................................... x List of Figures ............................................................................................................................................ xii List of Abbreviations ............................................................................................................................... xiv Acknowledgments .................................................................................................................................... xix Chapter 1. Introduction .............................................................................................................................. 1 1.1 Protein arginine N-Methyltransferases (PRMTs).................................................................................... 1 1.1.1 Arginine methylation ................................................................................................................... 1 1.1.2 PRMT structure and function ....................................................................................................... 3 1.1.3 PRMT substrate methylation activity .......................................................................................... 6 1.1.4 PRMT biological roles ................................................................................................................. 9 1.1.5 PRMT as therapeutic targets ...................................................................................................... 13 1.1.5 PRMT-PRMT interactions ......................................................................................................... 14 1.2 PRMT2 .................................................................................................................................................. 17 1.2.1 PRMT2 discovery ...................................................................................................................... 17 1.2.2 PRMT2 interacts with and regulates the activity of nuclear receptors ...................................... 17 1.2.3 Suspected PRMT2 methylation activity .................................................................................... 21 1.2.4 PRMT2 structure ........................................................................................................................ 27 1.2.5 PRMT2 proteomics investigations ............................................................................................ 34 1.2.6 PRMT2 heteromeric interactions .............................................................................................. 35 1.3 Hypothesis and research objectives ..................................................................................................... 38 Chapter 2. Protein arginine N-methyltransferase activity determination with filter binding and phosphor screening (FBAPS) assay ........................................................................................................ 39 2.1 Introduction ........................................................................................................................................... 39 2.2 Methods................................................................................................................................................. 41 2.2.1 Reagents ..................................................................................................................................... 41 2.2.2 Protein expression and purification ........................................................................................... 41 2.2.3 Enzyme assay ............................................................................................................................. 43 2.2.4 Scintillation counting ................................................................................................................. 43 viii 2.2.5 Densitometry and data analysis ................................................................................................. 44 2.2.6 SAM standards and unit conversions ......................................................................................... 44 2.2.7 KM and kcat determination ........................................................................................................... 44 2.2.8 IC50 determination ..................................................................................................................... 45 2.3 Results and Discussion ......................................................................................................................... 46 2.3.1 FBAPS assay conditions ............................................................................................................ 46 2.3.2 Enzyme kinetic determinations using the FBAPS assay............................................................ 48 2.3.3 Enzyme inhibition measured by the FBAPS assay .................................................................... 50 2.3.4 Concluding remarks ................................................................................................................... 52 2.3.5 PRMT2 and filter binding assays ............................................................................................... 54 Chapter 3. Development of PRMT2 binding methods ......................................................................... 56 3.1 Introduction ........................................................................................................................................... 56 3.2 Methods................................................................................................................................................. 58 3.2.1 Reagents ..................................................................................................................................... 58 3.2.2 Protein expression and purification ........................................................................................... 58 3.2.3 Differential scanning fluorimetry .............................................................................................. 58 3.2.4 Gel radiography ......................................................................................................................... 59 3.3 Results ................................................................................................................................................... 60 3.3.1 PRMT2 thermal stability characterization ................................................................................. 60 3.3.2 PRMT2 thermal stabilization by excipients ............................................................................... 62 3.3.3 PRMT2 response to small molecules ......................................................................................... 66 3.4 Discussion ............................................................................................................................................. 68 3.4.1 Human PRMT2 thermal melt and activity ................................................................................. 68 3.4.2 Exploring PRMT2 thermal stability with solution additives ..................................................... 69 3.4.3 PRMT2 interactions with PRMT inhibitors ............................................................................... 71 Chapter 4. Protein arginine N-methyltransferase 2 (PRMT2) plays a noncatalytic role in the histone methylation activity of PRMT1 ............................................................................................................... 73 4.1 Introduction ........................................................................................................................................... 73 4.2 Methods................................................................................................................................................. 75 4.2.1 Materials .................................................................................................................................... 75 4.2.2 DNA constructs .......................................................................................................................... 76 4.2.3 Protein expression and purification ........................................................................................... 77 4.2.4 His-tagged PRMT purification .................................................................................................. 77 4.2.5 GST-tagged PRMT purification ................................................................................................ 78 ix 4.2.6 Histone purification ................................................................................................................... 79 4.2.7 Differential scanning fluorimetry .............................................................................................. 80 4.2.8 Native PAGE ............................................................................................................................. 80 4.2.9 Histone methylation kinetics ...................................................................................................... 81 4.2.10 Gel radiography ....................................................................................................................... 81 4.3 Results ................................................................................................................................................... 82 4.3.1 PRMT2 ligand binding characterization ................................................................................... 82 4.3.2 SH3 domain effects on PRMT2 thermal stabilization .............................................................. 84 4.3.3 PRMT1/2 complex response to ligands ..................................................................................... 85 4.3.4 PRMT2 impact on PRMT1 substrate specificity ....................................................................... 90 4.4 Discussion ........................................................................................................................................... 102 4.4.1 PRMT heteromeric complexation ........................................................................................... 102 4.4.2 Human PRMT2 thermal stability characterization .................................................................. 103 4.4.3 Non-canonical SH3 domain functions ..................................................................................... 104 4.4.4 Requirements for PRMT1/2 complexation .............................................................................. 105 4.4.5 PRMT2 impact on substrate specificity ................................................................................... 105 4.4.6 PRMT2 and the histone code ................................................................................................... 106 4.4.7 Concluding remarks ................................................................................................................. 107 Chapter 5. Conclusions and future directions ...................................................................................... 108 5.1 Major thesis findings, significance, and impact .................................................................................. 108 5.1.1 FBAPS strengths and limitations ............................................................................................. 108 5.1.2 PRMT2 binding assays ............................................................................................................ 109 5.1.3 PRMT2 protein-protein interactions ........................................................................................ 110 5.2 Future directions ................................................................................................................................ 113 5.2.1 PRMT2 inhibitor design .......................................................................................................... 113 5.2.2 Interrogating the PRMT2 SH3 domain ................................................................................... 115 5.2.3 PRMT1/2 structural characterization ....................................................................................... 116 5.2.4 PRMT1/2 and epigenetic crosstalk .......................................................................................... 117 5.3 Conclusions ......................................................................................................................................... 118 References ................................................................................................................................................ 119 Appendix Figures .................................................................................................................................... 151 Appendix Tables...................................................................................................................................... 181 x List of Tables Table 1.1. Measures describing the structural alignment of PRMT1 and PRMT6 against PRMT2 using pairwise structural alignment ...................................................................................................................... 29 Table 2.1. Tabulated KM and kcat values comparing FBAPS against filter binding assay ........................... 50 Table 2.2. Tabulated IC50 and estimated KI values .................................................................................... 52 Table 4.1. Tabulated apparent kinetic parameters for PRMT1/2 methylation of histones H2A, H3, and H4 .................................................................................................................................................................... 98 Table 4.2. PRMT methylation activity towards calf thymus histones +/- PRMT2 ................................... 102 Table A1. Derived melting temperature (Tm) for PRMT2 from Figure 3.1C and D................................. 181 Table A2. One-way ANOVA statistical testing evaluating the melting temperature (Tm) from Figure 3.1C and D ......................................................................................................................................................... 182 Table A3. Derived melting temperature for PRMT2 (Tm1 and Tm2) from Figure 3.2A ............................ 183 Table A4. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.2A ........................................................................................................................................................... 184 Table A5. Derived melting temperature (Tm1 and Tm2) for PRMT2 from Figure 3.2B ............................ 185 Table A6. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.2B ........................................................................................................................................................... 186 Table A7. Derived melting temperature for PRMT2 (Tm1 and Tm2) from Figure 3.2C ............................ 187 Table A8. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure 3.2C ........................................................................................................................................................... 187 Table A9. Unpaired studentâs statistical testing evaluating the melting temperatures (Tm2) from Figure 3.2C ........................................................................................................................................................... 187 Table A10. Derived melting temperature for PRMT2 (Tm) from Figure 3.3A and B............................... 188 Table A11. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.3A and 3.3B ........................................................................................................................................... 189 Table A12. Derived melting temperature for PRMT2 (Tm) from Figure 3.4B, C, D ................................ 189 Table A13. Derived melting temperature for PRMT2 (Tm) from Figure 3.4A, E .................................... 190 Table A14. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.4 (for PRMT2 + SAM, Sinefungin, or Diamine 12) .................................................................................... 190 Table A15. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.4 (for PRMT2 + SAH, and/or MS023) ........................................................................................................ 190 Table A16. Derived melting temperature for PRMT2 (Tm1/Tm2) with ligands from Figure 4.1 ............... 191 Table A17. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure 4.1 ............................................................................................................................................................. 192 Table A18. Derived melting temperatures for PRMT2 (Tm1) from Figure A16 ....................................... 192 Table A19. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A16 ............................................................................................................................................................ 193 xi Table A20. Derived melting temperature for PRMT2H112Q (Tm1) from Figure A17 ............................. 193 Table A21. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A17 ............................................................................................................................................................ 193 Table A22. Derived melting temperature for PRMT2ÎSH3 (Tm1) from Figure A17 ............................... 194 Table A23. One-way ANOVA statistical testing evaluating the for PRMT2ÎSH3 melting temperatures (Tm) from Figure A17 ............................................................................................................................... 194 Table A24. Derived melting temperatures for PRMT2ÎSH3 (Tm) with H3 peptide from Figure A18 .... 195 Table A25. Derived melting temperature for PRMT1 (Tm) from Figure A19 .......................................... 195 Table A26. Derived melting temperature for PRMT1 (Tm) with increasing amount of H3 peptide from Figure A19 ................................................................................................................................................ 196 Table A27. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A19 ............................................................................................................................................................ 197 Table A28. Derived melting temperature for PRMT1/2 complex (Tm1 and Tm2) from Figure 4.2 ........... 197 Table A29. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2 from Figure 4.2 ..................................................................................................................... 198 Table A30. Derived melting temperature for PRMT1 and PRMT2ÎSH3 from Figure 4.2 ...................... 199 Table A31. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2ÎSH3 from Figure 4.2 ........................................................................................................... 200 Table A32. Derived melting temperatures for PRMT1 and PRMT2H112Q from Figure 4.2 .................. 200 Table A33. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2H112Q from Figure 4.2 ......................................................................................................... 201 Table A34. Derived melting temperature for PRMT1 and PRMT2 (at different concentrations) from Figure A19 ................................................................................................................................................ 202 Table A35. Tabulated data from densitometry analysis in Figure A26 .................................................... 206 Table A36. Tabulated data from densitometry analysis from Figure A27 ................................................ 207 Table A37. Tabulated data from densitometry analysis from Figure A28 ................................................ 207 Table A38. Tabulated data from densitometry analysis from Figure A29 ................................................ 208 Table A39. Relative densitometry analysis for Figure 4.5E ..................................................................... 208 xii List of Figures Figure 1.1. Protein arginine methylation reactions ...................................................................................... 2 Figure 1.2. PRMT1 structure ....................................................................................................................... 5 Figure 1.3. Globular domains of the mammalian PRMT family .................................................................. 7 Figure 1.4. Structural conserved homodimer conformation amongst PRMT enzymes .............................. 15 Figure 1.5. Structural alignment of PRMT1 and PRMT6 against PRMT2 using pairwise structural alignment .................................................................................................................................................... 28 Figure 1.6. Predicted structure of full-length PRMT2 ................................................................................ 30 Figure 1.7. PRMT2 binding interactions with SAH and Cp1 ..................................................................... 32 Figure 2.1. FBAPS assay layout and output ............................................................................................... 47 Figure 2.2. Quantitation of phosphor signal ............................................................................................... 48 Figure 2.3. Michaelis-Menten enzyme kinetics using the FBAPS assay .................................................... 49 Figure 2.4. Dose-dependent inhibition curves using the FBAPS assay ...................................................... 51 Figure 2.5. Measuring the linear range for PRMT2 methylation of RSF1 by filter binding assay ............. 55 Figure 3.1. PRMT2 thermal stability measured by differential scanning fluorimetry (DSF) ..................... 62 Figure 3.2. PRMT2 thermal stability as a function of osmolytes ............................................................... 64 Figure 3.3. PRMT2 thermal stability as a function of solution pH ............................................................. 65 Figure 3.4. PRMT2 stabilization by PRMT small molecule probes ........................................................... 67 Figure 4.1. Effects of ligand binding on PRMT2 thermal stability ............................................................ 83 Figure 4.2. PRMT1/2 response to ligands by DSF ..................................................................................... 87 Figure 4.3. Native PAGE gels of fluorescent PRMT1/2 heteromeric complexes ....................................... 89 Figure 4.4. PRMT1/2 methylation of calf thymus histones ........................................................................ 93 Figure 4.5. PRMT1/2 methylation of histones in different macromolecular contexts ................................ 96 Figure 4.6. PRMT2 influence on PRMT methylation of histones ............................................................ 101 Figure A1. PRMT2 splice variants ........................................................................................................... 151 Figure A2. PRMT2 STRING protein-protein interaction map ................................................................. 152 Figure A3. PRMT2 GeneMANIA protein-protein interaction map .......................................................... 153 Figure A4. Linear range determination using the FBAPS assay .............................................................. 154 Figure A5. Optimizing total number of washes required for the filter paper used for FBAPS assay ....... 154 Figure A6. Evaluating the effects of increasing protein load on radioactive detection ............................ 155 Figure A7. Evaluating the retention of methylated protein on P81 filter binding paper ........................... 156 Figure A8. PRMT2 purification analysis .................................................................................................. 157 Figure A9. CLUSTAL multiple PRMT sequence alignment by MUSCLE ............................................. 158 Figure A10. PRMT2 methylation of GST-GAR. ...................................................................................... 159 xiii Figure A11. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of osmolytes ............... 160 Figure A12. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of pH .......................... 161 Figure A13. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of small molecule probes .................................................................................................................................................................. 162 Figure A14. Purity of recombinant PRMT enzymes used in Chapter 4 ................................................... 163 Figure A15. Impact of ligands on PRMT2 thermal stability .................................................................... 164 Figure A16. PRMT2 bisubstrate thermal stabilization analysis ................................................................ 165 Figure A17. Thermal melt of PRMT2H112Q in DSF .............................................................................. 165 Figure A18. Thermal melt of PRMT2ÎSH3 in DSF ................................................................................ 166 Figure A19. Impact of H3 peptide on PRMT1 thermal stability .............................................................. 167 Figure A20. PRMT1/2 complex thermal stability ................................................................................... 168 Figure A21. PRMT1/PRMT2ÎSH3 complex thermal stability ............................................................... 169 Figure A22. PRMT1/PRMT2H112Q complex thermal stability ............................................................. 170 Figure A23. PRMT1/2 complex thermal stability varying protein concentrations ................................... 171 Figure A24. Impact of storage buffer on PRMT1 methylation of calf thymus histones .......................... 172 Figure A25. PRMT1E153Q/2 methylation of calf thymus histones ......................................................... 173 Figure A26. Gel radiography analysis of calf thymus histone methylation by PRMT1/2 ........................ 174 Figure A27. FBAPS analysis of H2A methylation by PRMT1/2 ............................................................. 175 Figure A28. FBAPS analysis of H3 methylation by PRMT1/2 ................................................................ 176 Figure A29. FBAPS analysis of H4 methylation by PRMT1/2 ................................................................ 177 Figure A30. FBAPS quantitation of individual histone methylation by PRMT2 ..................................... 178 Figure A31. Possible PRMT2 SH3 domain ligands ................................................................................. 179 Figure A32. AlphaFold structure of dimeric human PRMT2 ................................................................... 180 xiv List of Abbreviations aDMA â Asymmetric dimethylarginine AdOx â Adenosine dialdehyde AF â Activating function AR â Androgen receptor ATP â Adenosine triphosphate BiFC â Biomolecular fluorescence complementation BME â β-mercaptoethanol BRCA1 â Breast cancer gene 1 BRD â Bromodomain containing protein CARM1 â Coactivator-associated arginine methyltransferase 1 CDKN2B â Cyclin-dependent kinase inhibitor 2B CETSA â Cellular thermal shift assay COPR5 â Coordinator of PRMT5 CRC â Colorectal cancer Cryo-EM â Cryogenic electron microcopy CV â Column volume DDR â DNA damage response DGR â DSF-guided refolding DLS â Dynamic light scattering DMSO â Dimethyl sulfoxide DNMT â DNA methyltransferase DSB â Double strand breaks DSF â Differential scanning fluorimetry DPM â Disintegrations per minute DTT â Dithiothreitol xv EGF â Epidermal growth factor eGFP â Enhanced green fluorescent protein EGFR â Epidermal growth factor receptor EDTA â Ethylene diamine- N, N, Nâ², Nâ²- tetraacetic acid EGTA â Ethylene glycol-bis (β-aminoethyl ether)-N, N, Nâ², Nâ²-tetraacetic acid EMT â Epithelial-to-mesenchymal transition ER â Estrogen receptor EVH1 â Drosophila-enabled/vasodilator-stimulated phosphoprotein homology 1 domain FBAPS â Filter binding and phosphor screening assay FRET â Förster resonance energy transfer GAR â Glycine- and arginine-rich motif GFP â Green fluorescent protein GBM â Glioblastoma multiforme GraFix â Gradient fixation GST â Glutathione S-transferase HA â Hemagglutinin HCC â Hepatocellular carcinoma HDAC â Histone deacetylase HIV â Human immunodeficiency virus HMT1 â hnRNP methyltransferase 1 hnRNP â Heterogenous ribonucleoprotein IC50 â Half-maximal inhibitory concentration IKK â IκB kinase IMAC â Immobilized metal ion affinity chromatography IP â Immunoprecipitation IPTG â Isopropyl-β-D-1-thiogalactopyranoside xvi ITC â Isothermal titration calorimetry kcat âTurnover number KD â Dissociation constant KI â Inhibitor constant KM â Michaelis constant MAVS â Mitochondrial antiviral-signaling protein mCer â Monomeric cerulean mCit â Monomeric citrine mRNA â Messenger RNA MEF â Mouse embryonic fibroblast MLL1 â Mixed-lineage leukemia 1 MMA â Monomethylarginine MEP50 â Methylosome protein 50 MRN â Mre11/Rad50/Nbs1 complex MTA â Methylthioadenosine NF-κB â Nuclear factor κB NMR â nuclear magnetic resonance spectroscopy PAGE â Polyacrylamide gel electrophoresis PABP1 â Poly(A) binding protein 1 PCa â Prostate cancer PDGF â Platelet-derived growth factor PDGFR â PDGF receptor PGM â Proline-glycine-methionine PH â Pleckstrin homology domain PHD â Plant homeodomain PMSF â Phenylmethylsulphonyl fluoride xvii PRMT â Protein arginine N-methyltransferase RB â Retinoblastoma protein RBP â RNA-binding protein RCC â Renal cell carcinoma RMSD â Root mean square deviation RSF1 â Repressor splicing factor 1 SAH â S-adenosyl-L-homocysteine SAHH â SAH hydrolase SAM â S-adenosyl-L-methionine Sam68 â Src-associated in mitosis 68 kDa protein SAP â Spliceosome associated proteins SAXS â Small angle x-ray scattering sDMA â Symmetric dimethylarginine SEC â Size exclusion chromatography siRNA â Small interfering RNA SH3 â Src homology 3 domain shRNA â Short hairpin RNA SMAD â Suppressor of mothers against decapentaplegic homolog SMN â Survival motor neuron complex SPA â Scintillation proximity assay SPIN1 â Spindlin1 SPR â Surface plasmon resonance STAT â Signal transducer and activator of transcription SWI/SNF â Switch/Sucrose non-fermentable TDRD â Tudor domain TGF-β â Transforming growth factor β xviii TGFR â TGF-β receptor THW â Threonine-histidine-tryptophan TIM â Triose phosphate isomerase TLR â Toll-like receptor Tm â Melting temperature TNF-α â Tumor necrosis factor α TR â Thyroid hormone receptor UPLC-MS/MS â Ultra high-performance liquid chromatography with tandem mass spectrometry WH2 â WASP homology 2 domain 3R â Three non-consecutive arginine residues 53BP1 â Tumor suppressor p53 binding protein 1 xix Acknowledgments First and foremost, I would like to thank my supervisor, Dr. Adam Frankel, for taking me on as a graduate student and supporting me in all my endeavours through my graduate studies. Adam has been a constant source of enthusiasm and positivity throughout my tumultuous journey with PRMT2 and I could not have done this experience without his guidance and mentorship. In addition, I thank Adam for providing me with many opportunities to train the many students who came through the Frankel lab during my tenure, and for allowing me to represent the Frankel lab both locally and internationally. Adam has invested considerable time in helping me become a more effective scientific leader and communicator and I will always be grateful for this investment. Additionally, I would like to thank my committee members, Dr. Brent Page, Dr. Karla Williams, Dr. Filip van Petegem, Dr. Mark Harrisson, and Dr. Brian Rodrigues for their support and guidance through my graduate studies. My committee provided constructive criticism and helpful feedback that helped me attain my research and professional goals. I thank the past and present Frankel lab students for creating a positive, social, and stimulating working environment that greatly contributed to my graduate school experience. I owe a lot to my predecessor and mentor, Dr. Jennifer Brown, who trained me on all the lab techniques, shared her wisdom and knowledge of the PRMT field, and helped me become a better mentor. I thank the undergraduate students Diana Vasiliev, Jasmin Gill, Molly Benbow, and Rui Wen (Ivy) Liu for their valuable contributions to science in the Frankel lab and for being positive and pleasant people to work with. I thank the visiting international research scholars, Anke Gremmel, Verena Maïlander, and Cerissa van Walstijn for contributing their diverse scientific knowledge to various projects in the Frankel lab and for being patient and kind through foreign experiences. I thank MSc students Shireen Anbardan, Riley Prout-Holm, and Alana Hitsman for their dedication to the success of various Frankel lab projects, for sharing the highs and lows of research, and for being thoughtful and courteous human beings. I thank all the wonderful human beings who I have worked with in the Frankel lab for their friendship that made my experience as a graduate student a positive one. xx I thank to the many individuals who provided technical expertise and training for many of the techniques employed during my tenure as a graduate student. I thank Dr. Filip van Petegem and Dr. Steven Molinarolo for analyzing PRMT1/2 complexes with negative stain electron microscopy imaging and lending advice around characterizing protein complexes. I thank Dr. Natalie Strynadka and Helena Sverak for analyzing PRMT1/2 complexes by glycerol gradient ultracentrifugation and for thoughtful discussion around characterizing protein complexes. I thank Dr. Brent Page and Petar Iliev for providing technical and theoretical support with chemistry and thermodynamic screening techniques. I thank Dickson Lai and Dr. Lukas Hohenwarter for mass spectrometry training and support throughout my graduate studies. I thank Dr. Shyh-Dar Li and the Li lab for allowing me access to the dynamic light scattering instrument and training me with this technique. I thank Dr. Sandi Spencer Miko and her team in the proteomics division from the BC Cancer Research Centre (Genome Sciences Centre) in addition to Dr. Jason Rogalski from UBC Michael Smith Laboratories for proteomic analysis or various samples. I thank Jeff McCarthy from AdMare for access to their phosphor imager. I am very grateful for the funding and grant support from the University of British Columbia (the Shaughnessy Hospital Volunteer Society Fellowship in Health Care, the Elwyn Gregg Memorial Fellowship, the Douglas and Jean Bailey Scholarship, and the University of British Columbia Four-Year Doctoral Fellowship), from the Faculty of Pharmaceutical Sciences (the Santosh M. Wasan Graduate Student Travel Award in Pharmaceutical Sciences, and the John H. McNeill Scholarship) and from the Natural Sciences and Engineering Research Council of Canada (NSERC) (CGS-M and PGS-D) throughout the entirety of my graduate studies. I would like to thank my family and friends for their love, motivation, and encouragement that have allowed me to reach my personal and professional goals in life. A very special thanks to my parents, Mr. and Mrs. Glenn and Alana Rowley, and my brother, Mr. Nicholas Rowley, for their endless amounts of love, support, and guidance throughout life. My family raised me to believe in myself, think critically, and to be creative when pursuing my goals, and I would not be where I am today with them. xxi Last but certainly not least, I would like to thank my partner, Annette Kjell, for her unwavering emotional support, limitless patience, and for keeping me focused and on track throughout my graduate experience. 1 Chapter 1. Introduction 1.1 Protein Arginine N-Methyltransferases (PRMTs) 1.1.1 Arginine methylation Proteins are composed of different combinations of the twenty unique amino acids that are distinguishable by their distinct functional âRâ groups. Post-translational modifications (PTMs) are covalent attachments that are chemically appended to the functional âRâ groups of amino acids following protein synthesis that further enrich the chemical diversity of proteins. One common PTM on polypeptides is arginine methylation (1). Arginine is one of the twenty amino acids distinguishable by its basic guanidino group with the potential for five hydrogen bond donors and positive charge under physiological conditions (2). Frequently, arginineâs positive charge allows it to associate with anionic species through salt bridges, hydrogen bonding, and cation-pi interactions. Arginine can be methylated on one of its three guanidino nitrogen atoms (3). Methylation of the nitrogen atom replaces a proton with a single methyl group, thus abolishing one hydrogen bond donor whilst increasing hydrophobicity and steric size (2). The additional methyl group does not significantly alter the pKa of arginine (4). In eukaryotes, four different methylarginine species have currently been characterized: NÅ-monomethylarginine (ÅMMA), NÅ1, NÅ2-dimethylarginine (sDMA), NÅ1, NÅ1-dimethylarginine (aDMA) and Nδ-monomethylarginine (δ-MMA) (Figure 1.1; δ-MMA not shown in figure as it is not made via mammalian PRMT activity) (3). δ-MMA is generated by Rmt2p from yeast (no human homologue) but its unclear what purpose δ-MMA has (5, 6). ÅMMA, sDMA, and aDMA each have unique non-covalent interactions that dictate novel intermolecular interactions for the methylated polypeptide (7). 2 Figure 1.1. Protein arginine methylation reactions. (A) Chemical structures of SAM, S-adenosyl-L-methionine, the cofactor of PRMT enzymes, and SAH, S-adenosyl-L-homocysteine, the product of PRMT reactions. (B) PRMT methylation reactions in mammals. Methyl groups (green) transferred from SAM to terminal (η) nitrogen atoms on arginine residues. Abbreviations are as follows: SAM, S-adenosyl-L-methionine; SAH, S-adenosyl-L-homocysteine; NÅ-monomethylarginine (ÅMMA); NÅ1, NÅ2-dimethylarginine (sDMA); NÅ1, NÅ1-dimethylarginine (aDMA). The types of PRMT responsible for each reaction are indicated in red. 3 Arginine methylation can either promote or inhibit protein-protein interactions, depending on the type of methylarginine produced and which arginine is methylated (8). Typically, arginine residues are exposed on the surface of polypeptides and easily accessible to the methyltransferase enzymes responsible for methylation (9). As an example, the N-terminal region of histones do not have defined structure and are exposed in solution to enzymes, which is why the residues in this region are methylated. Specifically, core histones H2A, H3, and H4 are primary sites for arginine methylation (10). It has been found that the asymmetric dimethylation of H3R2 has an inhibitory effect on the binding of proteins to nearby amino acids (e.g., H3K4 methylation is inhibited as the lysine methyltransferase cannot bind to its substrate) (10). The addition of the methyl groups is enough to disrupt binding of any complexes that normally bind to H3K4me, a true testament to the impact of how a small modification can have such a profound impact (11). While some interactions are inhibited by arginine methylation, others are promoted. The Tudor domain (TDRD) is a canonical binder of methylated residues (arginine or lysine) (7). Tudor domains possess an aromatic cage that interacts with methyl arginine residue through specific electrostatic, hydrogen-bond, cation-pi, and Van der Waals interactions (12). The TDRDs responsible for binding methyllysine or methylarginine vary substantially in their aromatic binding cages. TDRD binding methyllysine tend to have a more shallow and broad aromatic binding pocket to accommodate the steric bulk of methyllysine (lysine can be bear up to three methyl groups), whereas TDRD binding methylarginine tends to have more narrow and deep aromatic binding pockets to accommodate the planar methylarginine (13, 14). Arginine methylation is required for assembly of the TDRD bearing spinal motor neuron (SMN) protein complex that regulates RNA splicing (13). Understanding how and why arginine residues are methylated has important implications in disease diagnostics and potential therapeutics. 1.1.2 PRMT structure and function Methyltransferase enzymes use S-adenosyl-L-methionine (SAM) to methylate a wide range of electron-rich substrates (8). SAM is a unique cofactor stored inside cells that methyltransferases use in nucleophilic, bimolecular SN2-like reactions due to its extremely labile methyl sulphonium group (Figure 1.1A) (15). Nucleophiles may attack this electrophilic methyl sulphonium group to yield methylated 4 substrate and S-adenosyl-L-homocysteine (SAH) (Figure 1.1A) (16). This methyl transfer is catalyzed by all SAM-dependent methyltransferases to which PRMTs belong (17). Structurally, PRMTs are part of the seven-β-strand superfamily of methyltransferases generally referred to as Class I methyltransferases (17). Class I methyltransferases are characterized by a Rossmann-like central fold and their ability to bind and methylate a wide array of substrates. Currently nine mammalian enzymes have been identified as PRMTs based on enzymatic activities, as well as sequence and structural similarities. The catalytic cores of all nine PRMTs possess an N-terminal Rossmann-like catalytic domain and a C-terminal domain containing a beta barrel and dimerization arm (Figure 1.2A) (18). Within the N-terminal catalytic domain reside three alpha helices (αX, αY, αZ) that form a lid over two conserved motifs within the active site: the double glutamic acid loop (the double-E-loop) (18), and the threonine-histidine-tryptophan (THW) loop (extending from the β-barrel domain to contact the SAM binding site) (Figure 1.2B) (17). These helices serve two additional functions: 1) forming a landing pad for the dimerization arm of other PRMT monomers to form catalytically active dimer complexes, and 2) providing substrate specificity within a peptide binding groove (19â21). In PRMTs, the peptide binding groove is only formed following the binding of SAM in the Rossmann-like binding domain (22). SAM binding causes a PRMT conformational change that enables formation of the peptide binding groove and allows catalytically active dimers to form (22â25). PRMTs will then methylate the peptide substrate either once or twice (in a processive manner) (22), depending on the type of PRMT. 5 Figure 1.2. PRMT1 structure. (A) X-ray crystal structure of PRMT1 (PDB ID: 6NT2) colored according to secondary structure elements (beta strand, blue; beta turn, green; random coil, grey; alpha helix, red). (B) Zoomed-in view of the double-E-loop of PRMT1 (PDB ID: 1OR8) showing conserved residues that form double-E and THW loops (with carbon atoms in yellow). The structure was solved with a peptidic arginine residue (carbon atoms in purple) bound between the double-E-loop. AaXaYaZaAaBBH293E144E153W294T292PeptideArginine6 The nine PRMTs can be divided based on the type of methylarginine they produce: MMA, aDMA, or sDMA (8). Type I PRMTs (PRMT1, PRMT2, PRMT3, PRMT4 (a.k.a. coactivator-associated arginine methyltransferase 1, CARM1), PRMT6, and PRMT8) make MMA and aDMA; Type II PRMTs (PRMT5 and PRMT9) make MMA and sDMA; Type III PRMTs (PRMT7) make only MMA (Figure 1.1B) (17). Recent studies have revealed that steric and hydrogen bonding networks around the double E loop dictate enzymatic activity (26). Two glutamic acid residues that make up part of the double E loop and one histidine residue from the THW motif perform two tasks: coordinating the methyl sulphonium group for methyl transfer through salt bridges and hydrogen bonding (18), and deprotonating the guanidino group to increase its nucleophilicity (Figure 1.2B) (27). Type III PRMTs have limited space around the double E loop and can only facilitate a single methyl transfer about the NÅ1 guanidino atom, whereas Type I and II PRMTs have an extra cavity to facilitate the transfer of an additional methyl group (26). Type II PRMTs perform symmetric dimethylation on the NÅ1 atom since the NÅ2 atom is restricted by hydrogen bonding in the double E loop. The same hydrogen bond network is not observed in Type I PRMTs (26). The peptide specificity conferred to each of the nine distinct PRMTs stems from its unique peptide binding groove (28). The peptide binding groove is defined as a long and hydrophobic channel that stretches across the enzyme face (29). It is the structural features of this channel, such as acidity to interact with positive charges, that defines the peptide binding specificities. In addition to the conserved Rossmann-like fold and β-barrel domain, certain PRMTs possess unique domains involved with substrate binding or cellular localization (Figure 1.3). 7 Figure 1.3. Globular domains of the mammalian PRMT family. Abbreviations are as follows: EVH1, Drosophila-enabled/vasodilator-stimulated phosphoprotein homology 1; SH3, Src homology 3; TIM, triose isomerase phosphate; TPR, tetratricopeptide repeats; ZnF, zinc-finger. PRMT2 contains an N-terminal Src homology 3 (SH3) domain that can bind proline-rich motifs and has been implicated in protein-protein interactions with potential substrates (30). PRMT3 contains an N-terminal zinc-finger domain that coordinates zinc ions to facilitate interactions with RNA associated proteins (31). PRMT4 contains an N-terminal PH (pleckstrin homology)-like domain that adopts a similar fold to Drosophila-enabled/vasodilator-stimulated phosphoprotein homology 1 domain (EVH1) and has been implicated in binding proline-rich motifs (32). PRMT5 contains a large N-terminal triose phosphate isomerase (TIM) barrel domain that forms a heteromeric complex with methylosome protein 50 (MEP50) (33). The PRMT5-TIM barrel interaction with MEP50 results in the formation of a large hetero-octameric complex with molecular weight ~450 kDa (33, 34). Unlike the other PRMT enzymes that do not require binding partners, the methylation activity of PRMT5 is maximized upon formation of a heteromeric complex with MEP50 (34). While the N-terminal auxiliary domains of PRMT2, 3, 4, and 5 are independently folded, it has been observed that removal of these domains can decrease or inhibit catalytic 8 activity of their associated PRMT enzyme (31â33, 35). PRMT7 contains two catalytic methyltransferase domains (likely through gene duplication) separated by a zinc-finger motif, yet only the N-terminal methyltransferase domain, and not the C-terminal methyltransferase domain, can functionally bind SAM (36). The active N-terminal methyltransferase domain of PRMT7 can interact with the inactive C-terminal methyltransferase domain to form a pseudo-homodimer (37). PRMT8, while sharing >80% sequence similarity to PRMT1, is N-terminally myristoylated in cells, making it the only PRMT enzyme that localizes exclusively to the plasma membrane (38, 39). Like PRMT7, PRMT9 also contains a duplication of the methyltransferase domain and is thought to exist as a pseudo-homodimer, but it also contains three tandem tetratricopeptide repeats (TPR) thought to be important for protein-protein interactions (40, 41). PRMT1 and PRMT6 do not contain any auxiliary domains (42, 43). 1.1.3 PRMT substrate methylation activity PRMTs exhibit broad substrate specificities and catalytic outputs. PRMT1 is the primary Type I PRMT in mammalian cells, responsible for ~85% of all arginine methylation (44). While PRMT1 displays broad substrate specificities, it has a clear preference for arginine residues flanked by one or more glycine residues referred to as the glycine- and arginine-rich (GAR) motif (3). Proteins bearing the GAR motif are frequently associated with binding nucleic acids, given the cationic arginine forms electrostatic interactions with the negative backbone of DNA/RNA (45). Proteomic investigations found that most proteins bearing aDMA marks belong to the heterogeneous ribonuclear protein (hnRNP) family, involved with pre-mRNA splicing and RNA shuttling (46). While PRMT1 is responsible for generating the majority of the aDMA in cells, its catalytic turnover number kcat, is in the same order of magnitude as PRMT3, CARM1, PRMT5, and PRMT6 (47). While CARM1 and PRMT6 can methylate the GAR motif to varying degrees, they have distinct substrate methylation preferences. CARM1 preferentially methylates arginine residues adjacent to proline-glycine-methionine (PGM) motifs (48). Similar to the RGG motif, PGM motifs are frequently associated with splicing factors and RNA binding proteins (RBPs) (49). PRMT6 and PRMT7 both share the same preferred methylation motif of RXR, where X is any amino acid (3, 50). PRMT5 is the major Type II methyltransferase responsible for the majority of 9 sDMA production in cells with a wide range of substrates, but no distinct methylation preference. A study into PRMT5 in vivo methylation of histones found that PRMT5 substrate specificity could change in the presence of the interacting protein, coactivator of PRMT5 (COPR5) (51). PRMT9 is the only other Type II PRMT, that has been shown to methylate the splicing factors, SF3B2 and SAP145, and the mitochondrial antiviral signaling protein (MAVS), but has no other reported methylation activity (40, 41, 52). There has been no enzyme kinetics study conducted on PRMT9 with its substrates in vitro, therefore the kcat has yet to be quantified, but detectable methylation signal was obtained after a one-hour incubation, which is comparable with the more active PRMTs (52). PRMT8 shares greater than 80% sequence similarity with PRMT1 and are both capable of methylating the GAR motif, but the kcat for PRMT8 is an order of magnitude lower than PRMT1 (53). PRMT2 by far has the lowest catalytic activity output of all the reported PRMT enzymes, with a kcat 800-fold lower than PRMT1 towards proteins with a GAR motif (19). 1.1.4 PRMT biological roles While hnRNPs and pre-mRNA processing proteins contain the majority of the methylated arginine residues in mammalian cells, proteins involved with chromatin remodeling, transcription, splicing, translation, endosomal trafficking, and cytoskeletal rearrangement have also been identified as PRMT substrates (10, 54â57). PRMTs mediate a wide range of cellular functions either through protein-protein interactions or through the methylation of substrates (3). Homozygous deletion (or knock-out) of PRMT1 and PRMT5 in mice results in embryonic lethality, whereas CARM1 knock-out mice died shortly after birth, demonstrating that arginine methylation is essential for survival (2, 8). The major biological roles of the pre-dominant PRMT enzymes (PRMT1, CARM1, PRMT5, PRMT6) contributing to cell survival are reviewed below. 10 Splicing regulation Proteins bearing the GAR motif are found in RBPs that are associated with RNA shuttling (between the nucleus and cytoplasm) and pre-mRNA splicing (58). PRMT5:MEP50 serves a direct role in splicing regulation through methylation of Sm proteins that are recognized by the Tudor domain of the SMN complex to promote small nuclear ribonucleoprotein (snRNP) maturation (13, 59). The dysregulation of PRMT5:MEP50 activity towards Sm proteins can lead to reduced assembly of spliceosome complexes and retention of introns (60). CARM1 and PRMT9 also methylate spliceosome associated proteins (SAP) that enhance the assembly of spliceosome complexes (SAP49 and SAP145) (41, 48). PRMT1 can methylate a broad range of RBPs associated with pre-mRNA splicing to alter their subcellular localization and activities (61â64). It was found that PRMT1 methylation of Src associated during mitosis 68 kDa (Sam68) can directly inhibit the ability of Sam68 to bind to poly (U) RNA and transport it from the nucleus (65). PRMT1 has also been shown to methylate hnRNPA1, a protein involved with mRNA alternative splicing, which perturbs its interaction with splicing related proteins (62, 66). DNA damage repair pathway PRMT1 plays a vital role in the DNA damage repair (DDR) pathway through arginine methylation of the GAR motif in MRE11, the exonuclease component of the MRE11/RAD50/NBS1 (MRN) complex responsible for repairing DNA double-strand breaks (DSBs) (67). Arginine methylation of the GAR motif of MRE11 is required for functional exonuclease activity. PRMT1 also methylates tumor suppressor P53 binding protein 1 (53BP1) and breast cancer gene 1 (BRCA1), other proteins involved in regulating DDR response (68, 69). PRMT5 has also been shown to can methylate proteins involved in DDR pathways (70â73). The dysregulation of PRMT1 or PRMT5 can lead to impaired DDR. compromised genomic integrity, and has been associated with cancer (74). 11 Cell signaling PRMT1 and PRMT5 are both involved in regulating how cells respond to growth-factor stimulus through cell-surface transmembrane receptor signaling pathways (58). A major signaling pathway frequently dysregulated in cancer progression is the epithelial-mesenchymal transition (EMT) that promotes cellular differentiation, invasion, and apoptosis resistance (75). The cell signaling pathways controlling EMT are frequently upregulated in actively metastasizing cancer including, non-small cell lung cancer and breast cancer (75â78). EMT signaling pathways are regulated by the cytokine transforming growth factor β (TGF-β) that binds transforming growth factor β receptor (TGFR) and trigger the activation of suppressor of mothers against decapentaplegic homolog (SMAD) proteins that can control genes related to cellular differentiation (79). PRMT1 is responsible for the methylation of the inhibitory SMAD6 and SMAD7, to promote their dissociation from TGFR, that allows the activation of SMAD3 to express proteins facilitating the EMT (80). Arginine methylated SMAD6 associates with MyD88, an adaptor protein required for the response of toll-like receptors (TLR), and inhibits TLR signaling pathways (81). TLR activation is linked to nuclear factor κB (NF-κB) signaling that regulate genes associated with inflammatory or immune response, that are frequently inhibited in cancer. PRMT5 can methylate the p65 subunit of NF-κB, an activating mark that promotes its transcriptional activity (82). PRMT1 and CARM1 have also been shown to serve as transcriptional coactivators of NF-κB (83, 84). PRMT5 also functions as a repressor of the epithelial growth factor (EGF) and promoter of the platelet derived growth factor (PDGF) signaling pathways through methylation of the receptors or downstream effectors (85, 86). Transcriptional regulation PRMTs have been identified as transcriptional coactivators, attributed to their propensity to methylate histones, transcription factors, and nuclear receptors in addition to forming protein-protein interactions with nuclear receptors and/or transcriptional coactivators (87â90). CARM1 was the first PRMT shown to be capable of regulating gene transcription by directly interacting with the transcriptional coactivator GRIP1 to enhance the transcription of the androgen receptor (AR), estrogen receptor (ER), 12 and thyroid hormone receptor (TR) (using transcriptional reporter assays) (87). PRMT1 was also identified as an interaction partner for GRIP1 and SRC-1 coactivators, through Glutathione S-transferase affinity tag (GST)-pulldown, and further showed a synergistic enhancement of transcriptional activity of AR, ER and TR (89). PRMT1 and CARM1 were demonstrated to have high activity against histone H4 or histone H3, respectively, sharing only histone H2A as a substrate, but not a preferred substrate for either enzyme (87, 91). This suggested to the authors that the coactivator properties of PRMT1 and CARM1 were unique from their histone methyltransferase activity. PRMT5 and PRMT6 have also been shown to interact with and regulate the transcriptional activities of transcriptional repressors and activators, respectively (92â95). Nuclear receptors and transcriptional machinery are recruited to gene promoters through protein-protein interactions with nucleosomes and these interactions are highly dependent on the PTMs decorating histone N-terminal tails and surface residues. Arginine methylation of histones are an important component of the âhistone codeâ, a term used to describe the post-translational modification of histone proteins (96). Histone PTMs are epigenetic modifications since they serve important regulatory roles in gene transcription without directly impacting the genetic code. The location and type of methylarginine deposited on histone N-terminal tails are strictly regulated since they can either promote or restrict access to the genes wrapped in the nucleosomes. PRMT1 and CARM1 are responsible for the formation of H4R3me2a, and H3R17me2a/H3R26me2a, respectively, which are associated with active gene promoters, whereas PRMT5 and PRMT6 are responsible for the formation of H4R3me2s/H3R8me2s or H3R2me2a, respectively, associated with silenced gene promoters (10, 91, 97â100). H4R3me2a and H3R17me2a marks can be read by TDRD3, a transcriptional coactivator, to promote transcription (101). The H3R26me2a mark has not been associated with any specific effector protein involved with transcriptional regulation, but it has been associated with the H3K27Ac (acetylation) (102). Histone tails bearing the H4R3me2s mark recruit the de novo DNA methyltransferase (DNMT3A) through its plant homeodomain (PHD) domain, to methylate gene promoters or recruit histone deacetylase (HDAC) enzymes, both resulting in reduced gene expression (100, 103, 104). There is 13 crosstalk between the H3R8me2s mark and other histone marks on histone H3, such as the prior acetylation of either H3K9 or H3K14 prevents PRMT5 methylation of H3R8, while the deposition of H3R8me2s prevents the methylation of H3K9 by G9a (105, 106). H3R2me2a exerts its influence on the epigenetic code through preventing the binding of effector molecules to the H3K4me3 mark in addition to preventing the MLL1 complex from methylating H3K4 (11, 107). Dysregulation of PRMT1 and CARM1 histone arginine methylation is associated with cancer programming through enhanced transcriptional activation of oncogenes and uncontrollable cell proliferation (108â115). PRMT5 and PRMT6 are also associated with oncogenic reprogramming, but through the deposition of transcriptional repressive marks on tumor suppressor genes (11, 97, 98, 106, 107, 116â119). 1.1.5 PRMTs as therapeutic targets Arginine methylation is a crucial PTM in the regulation of protein-protein interactions that are important for cellular signaling pathways associated with correct physiological functions (3, 8). Dysregulation of PRMT expression can result in disruption of multiple cellular signaling pathways and lead to the development of disorders including, neurodegenerative, cardiovascular, and neuromuscular disease, and cancer (3, 120â123). In disease pathologies characterized by PRMT overexpression, PRMTs present themselves as therapeutic targets where inhibition of their catalytic activity will reduce cellular arginine methylation to attenuate dysregulated signaling pathways (2, 124, 125). There have been significant efforts put forth in the development of targeted small molecule inhibitors against PRMT activity (126, 127). These molecules either target the SAM binding domain, the peptide-binding domain, or both (defined as bisubstrate inhibitors). Small molecule inhibitors exclusively targeting the SAM binding domain serve as broad-spectrum methyltransferase (or Pan-MTase) inhibitors since they can bind indiscriminately to any protein with a SAM binding domain (Rossmann fold) and have limited therapeutic value (128). Examples of Pan-MTase inhibitors include SAH, sinefungin, and methylthioadenosine (MTA), that share the adenosyl-L-methionine like scaffold and serve as substrate competitive inhibitors (128â130). Specificity against PRMTs can be achieved by targeting small molecule inhibitors against the peptide-binding groove. A common chemical scaffold used to target the peptide 14 binding groove of PRMT enzymes is the extended alkyl arylamine, which is present in the small molecule inhibitors MS023, EPZ020411, and Diamine 12 (131â133) (for a more comprehensive review of PRMT small molecule inhibitors, refer to: (126)). With each of these molecules, a terminal methyl ethylene-diamine serves as a guanidine mimic whereas the remainder of the molecule can be conjugated with varying functional groups to specifically target the desired PRMT of interest (131). There has been some success in generating peptide-based bisubstrate inhibitors against PRMTs by chemically appending a SAM-like molecule to an arginine residue within the peptide (134, 135). These inhibitors not only bind to both SAM and peptide binding grooves, but further serve as transition state analogues (mimicking the pentavalent trigonal bipyramidal transition state of arginine methylation reactions) (136). With peptide based PRMT inhibitors, specificity is achieved by using peptides specific to the PRMT of interest like H4 for PRMT1 or poly(A) binding protein 1 (PABP1) for CARM1. Several PRMT inhibitors have made it into human clinical trials as cancer therapeutics (137â140). So far, only inhibitors against PRMT1 and PRMT5 have entered clinical trials. Developing a stronger understanding of the biochemistry and interactions of other PRMTs may lead to the development of more targeted inhibitors and potentially future disease therapeutics. 1.1.6 PRMT-PRMT interactions PRMT enzymes are ubiquitously expressed amongst different tissues (except PRMT8, which is expressed pre-dominantly in the brain) and found in both the nucleus and cytoplasm with the opportunity of interacting with a range of cellular proteins, including other PRMTs (8). The crystal structures of PRMT1-8 demonstrate highly similar homodimers (or pseudo-homodimers) mediated through the canonical dimer arm and helical docking pad (except PRMT5 where the dimerization arm interacts with the TIM barrel domain instead of the helical docking pad) (Figure 1.4) (20, 36, 38, 141â144). Molecular dynamics stimulations of PRMT1 reveal the dimer arm and the three alpha-helices forming the âdocking padâ for the dimerization arm are highly mobile (4). The N-terminal domains of PRMTs also have a high degree of flexibility, which has been demonstrated by the CARM1 crystal structures where only in the presence of the cofactor, SAH, are the N-terminal residues resolved (143, 145, 146). The crystal 15 structures for the majority of PRMTs do not show N-terminal residues, attributed to their high flexibility, which could imply these regions can participate in protein-protein interactions. There is a high degree of similarity between all PRMT enzymes amongst the residues in the PRMT dimer interface, providing the possibility that PRMTs could form heteromeric complexes with one another, potentially with novel substrate specificities (19). Figure 1.4. Structural conserved homodimer conformation amongst PRMT enzymes. Images rendered using BIOVIA Discovery Studio 2021. Each homodimer is composed of two identical monomers, distinguished by different colors. The first evidence of a heteromeric PRMT interaction accompanied the identification of PRMT3, where the authors used PRMT1 as a bait protein in a yeast two-hybrid screen and identify PRMT3 as an interaction partner (147). While no follow-up study further characterized the PRMT1-PRMT3 hetero-oligomer in humans, another group identified that PRMT1 and PRMT3 (from Trypanosoma brucei) form PRMT1:SAHPDB ID: 6NT2HomodimerPRMT2:SAHPDB ID: 5FULHomodimerCARM1:SAHPDB ID: 2V74HomodimerPRMT6:SFGPDB ID: 4C03HomodimerPRMT7:SAHPDB ID: 4C4APseudo-homodimerPRMT8:SAHPDB ID: 5DSTHomodimer16 an obligate heteromeric tetramer where the catalytically active subunit, PRMT1, only has activity when complexed with PRMT3 (148). Several studies have also corroborated an interaction between PRMT1 and PRMT8 through yeast two-hybrid screening, co-immunoprecipitation (co-IP) and mass spectrometry studies of the human interactome (53, 149â151). An in vitro PRMT1-PRMT6 heterodimeric interaction was identified by performing Förster resonance energy transfer (FRET) experiments with monomeric cerulean (mCer) fused to PRMT1 and monomeric citrine (mCit) fused to PRMT6 (23). The authors noted that the concentration of enzymes required to yield FRET signal from PRMT1-PRMT6 was far greater than the KD of dimerization for homodimerization for either PRMT1 or PRMT6, indicating the heteromeric PRMT1-PRMT6 dimers unlikely form an appreciably large population in cells (23). It was also recently identified that PRMT1 can methylate PRMT6 to decrease its activity against histone H3 (152). PRMT6 was also recently demonstrated to interact directly with PRMT5 to promote the epigenetic silencing of cyclin-dependent kinase inhibitor 2B (CDKN2B), which in turn promotes colorectal cancer progression (CRC) (153). PRMT2 has also been implicated in heteromeric protein interactions with other PRMTs. PRMT2 is one of the most poorly characterized PRMT enzymes, largely attributed to its low in vitro activity that makes it difficult to study (35). PRMT2 has been implicated in many of the same pathways as the more active PRMTs, such as pre-mRNA splicing, cellular signaling, transcriptional regulation, and epigenetic modifications. Several studies have emerged demonstrating a specific protein-protein interaction between PRMT2 with PRMT1, CARM1, PRMT6, and PRMT8 (19, 39, 154, 155). Unlike the other heteromeric PRMT interactions described previously, PRMT2 interaction with PRMT1 augmented the activity of PRMT1 towards histone H4 in vitro and increased global arginine methylation in cells when overexpressed with PRMT1 (19). The influence of PRMT2 on the substrate specificity, cellular localization, or protein stability of the other PRMT enzymes is unknown. It is plausible that PRMT2 could form a ternary complex with these proteins and substrates listed previously, to regulate their activities. The biological role of PRMT2 may not be through its methylation activity, but instead it 17 may be to regulate the activities of other enzymes. An analysis of the literature surrounding PRMT2 interactions does not preclude this hypothesis. 1.2 PRMT2 1.2.1 PRMT2 discovery PRMT2 (initially termed HRMT1L1) was discovered by two independent groups in 1997-1998 based on homology to rat PRMT1 and yeast human methyltransferase 1 (Hmt1) (43, 156). Early genetic analysis of PRMT2 identified a distinguishable N-terminal SH3 domain, and a methyltransferase domain with 60% similarity to yeast HMT1 (156). Using Northern blot analysis, PRMT2 RNA was found to be expressed to variable degrees in various adult and fetal human tissues (156). Silver and coworkers were unsuccessful in characterizing in vitro enzymatic activity from GST-PRMT2 or GST-PRMT2ÎSH3 (a PRMT2 truncation lacking the SH3 domain) against the yeast Npl3 hnRNP protein or human hnRNPA1 (using a radiometric methylation assay) (156). These results were complemented by a sophisticated yeast-based methylation assay where cell growth at non-permissive temperatures is dependent on overexpression of an active arginine methyltransferase (PRMT1 or Hmt1) to overcome the temperature-sensitive mutant npl3-1 (A254V) to facilitate export of poly (A)+ RNA from the nucleus to cytoplasm (157). Using this temperature- and methylation-sensitive yeast assay, PRMT2 could not rescue the synthetic lethal phenotype of Îhmt1 npl3-1 yeast at the elevated temperature. The authors provide several explanations for the lack of activity from PRMT2, including the potential need for an interaction partner for the SH3 domain of PRMT2, the hnRNP proteins evaluated may not be the ideal substrate for PRMT2, or that PRMT2 may no longer serve as an active methyltransferase (156). 1.2.2 PRMT2 interacts with and regulates the activity of nuclear receptors Shortly after the characterization of PRMT1 and CARM1 as secondary coactivators of the p160 family of coactivators, other groups started to investigate whether other PRMTs had coactivating properties (87, 89). Using yeast-two hybrid screening, GST-pulldown assays, and luciferase reporter assays, Qi and coworkers identified that PRMT2 could directly bind and enhance the transcriptional activity of the estrogen receptor α (ERα). Using truncations of both the ERα and PRMT2, the authors 18 provide evidence that the PRMT2 methyltransferase domain (amino acids 133-275) bound to the transcriptional activation function domain 1 (AF-1) and the DNA-binding domain of ERα (90). It was also demonstrated that PRMT2 can bind tritiated SAM (3H-SAM) using a filter-binding assay that retains protein bound to SAM while washing away unbound cofactor. The authors performed site-directed mutagenesis on motif I of the PRMT2 Rossmann-like fold (ILDVGCGTG) according to a previous study that reported impaired SAM binding by mutating the two underlined glycine residues to arginine (158). There were no crystal structures of Hmt1, PRMT1, or PRMT2 at this point and the mutation was selected based on conservation of these glycine residues in motif I amongst other methyltransferase enzymes (159). The G-to-R mutations (G145R, G147R) drastically reduced the ability of PRMT2 to bind SAM, and unlike the PRMT2ÎSH3 mutation, was shown to impair PRMT2 coactivator function using an ER transactivation assay. Another significant finding in this study was that GST-PRMT2 could self-interact via GST-pulldown assay, demonstrating the capacity of PRMT2 to form homodimers or homooligomers (90). This study demonstrated that the PRMT2 methyltransferase domain was essential for its ability to interact with and activate the estrogen receptor but did not identify the estrogen receptor as a PRMT2 substrate. These data demonstrate that PRMT2 regulates transcription through its interactions with the estrogen receptor. The association between PRMT2 and ERα has implicated PRMT2 and PRMT2 variants with breast cancer (160â164). Zhong and coworkers identified elevated PRMT2 mRNA expression in breast cancer tissues compared to normal tissues (160, 161). This group further identified several unique PRMT2-splice variants that contain intact N-terminal SH3 and SAM-binding domains but lacking segments of the β-barrel domain (Figure A1) (160, 161). These PRMT2 splice variants were all capable of binding to and improving the transactivation of ERα, as demonstrated by co-IP and ERα-dependent luciferase reporter assay (160, 161). Subcellular localization experiments (using transfected GFP-fusion PRMT2 splice variants into MCF7 cells) demonstrated that PRMT2 variants possessing the dimerization arm localized predominantly to the nucleus, whereas variants lacking this dimerization arm localized in the cytoplasm and nucleus (160). PRMT2 and PRMT2β were further demonstrated to suppress the growth 19 and division properties of MCF7 cells, through reducing the binding affinity of ERα to the AP-1 domain in the CCND1 promoter, yielding concomitant reduction of cyclin D1 expression (162). Oh and coworkers linked decreased PRMT2 expression in breast cancer with decreased cyclin D1 production, in contrast to what was identified by Zhong and coworkers (163). Oh and coworkers further demonstrated that genes upregulated as a function of PRMT2 downregulation contribute to increased survival rates whereas the genes upregulated as a function of PRMT2 upregulation negligibly impacted survival rates (in breast cancer patients) (163). These contradictory findings demonstrate that the exact mechanisms of PRMT2 in breast cancer remains largely enigmatic and cannot be reduced to its association with the estrogen receptor. The initial discovery of PRMT2 interactions with nuclear proteins (hnRNPs, nuclear receptors, and coactivators) led many researchers to inquire whether PRMT2 could interact with other proteins. In 2006, Ganesh and coworkers identified that PRMT2 negatively regulates the NF-κB signaling pathway (165). The NF-κB transcription factor regulates the expression of genes involved in immune and inflammatory response, but it is sequestered in the cytoplasm by the IκB protein complex to prevent gene expression (166). The IκB complex can be phosphorylated by inducible IκB kinase 1 or 2 (IKK1/IKK2), triggering it to release NF-κB heterodimers (p50/p65) that can translocate to the nucleus, bind κB-DNA elements, recruit transcriptional machinery, and promote gene expression (166). Using κB-reporter assays, it was demonstrated that PRMT2 completely inhibited NF-κB-dependent transcription (165). Several PRMT2 mutants were generated including, a mutant of the first four amino acids in the first motif of the PRMT2 Rossmann-like fold (ILDVGCGTG) to alanine, a PRMT2 splice variant lacking the C-terminal β-barrel, and a PRMT2 truncation containing only the SH3 domain (165). The PRMT2 methyltransferase domain was found to be indispensable for inhibiting NF-κB transcription, whereas mutations impairing SAM binding had no effect on the PRMT2-based NF-κB transcriptional inhibition. It was reported that recombinant-PRMT2 expressed and purified from either bacteria or mammalian sources were incapable of methylating histones, p65, p50, IκB, or hnRNP U (165). The authors found that the C-terminal β-barrel domain and, to a lesser extent, the methyltransferase domain of PRMT2 could directly 20 interact in vivo with the ankyrin repeats of IκBα (one of the constituents of the IκB complex) using co-IP (165). The authors proposed that PRMT2 inhibits NF-κB transcription through binding IκBα (that is associated with p50/p65 heterodimers), preventing the NF-κB/IκB complex from binding to DNA, and further preventing its nuclear export. PRMT2 inhibits NF-κB induced transcription, making the cells more susceptible to apoptosis without expression of the inflammatory response proteins encoded by NF-κB (165). The authors did not evaluate if PRMT2 inhibition of NF-κB signaling is related to PRMT5 methylation of NF-κB or the transcriptional activation of NF-κB by either PRMT1 or PRMT5 (82, 83). Concomitantly with the characterization of a PRMT2 NF-κB interaction, the Nabel laboratory characterized a PRMT2 ternary complex with the retinoblastoma gene product (RB) and the E2F transcription factor to regulate control of the cell cycle (167). The Nabel laboratory demonstrated that RB is selectively coprecipitated by PRMT2 (not PRMT1, 3, or CARM1), and the interaction is primarily between the PRMT2 SAM binding domain and the RB transactivation domain. It was further demonstrated that PRMT2 could coprecipitate E2F in the presence of RB, and this association inhibited E2F transcriptional activity using E2F reporter assays (167). In this study, PRMT2 null mice reported higher E2F activity than wild-type mice, and PRMT2-/- mouse embryo fibroblasts (MEFs) demonstrated earlier S-phase entry, consistent with the hypothesis that PRMT2 inhibits E2F activity (165). The authors postulated that in addition to suppressing the transactivation domain of RB, PRMT2 may recruit other co-repressors or impact the activity of other co-repressors associated with RB, including histone deacetylases (HDAC), Switch/Sucrose non-fermentable (SWI/SNF) chromatin remodeling complex, CtBPâpolycomb complex, lysine methyltransferase (Suv39H1), and DNMT1 (168â172). While the authors did not demonstrate that E2F is a PRMT2 methylation substrate, other groups have demonstrated that E2F asymmetric dimethylation by PRMT1 leads to DNA damage-induced apoptosis, whereas symmetric dimethylation by PRMT5 antagonizes this effect (173). There have been no studies evaluating whether PRMT2 is involved in the PRMT1 or PRMT5 methylation of E2F. The androgen receptor (AR) was another nuclear receptor identified as a PRMT2 interaction partner (174). An interaction between the AR and PRMT2 was identified by yeast two-hybrid screening 21 and validated through GST-affinity pulldown experiments that showed the interaction was AR ligand-dependent and mediated through both the PRMT2 SAM-binding and SH3 domains (174). Using AR reporter assays in cells, it was demonstrated that the determinants identified for the PRMT2-AR interaction (AR ligand, PRMT2 SAM-binding and SH3 domains) were also required for PRMT2 to coactivate the AR. Interestingly, the PRMT2 coactivation of the AR was inhibited with increasing amounts of SAH (174). PRMT2 failed to show synergistic activation of the AR in presence of other coactivators, CARM1 or GRIP1. AR signaling is frequently dysregulated in prostate cancer (PCa) and based on the interaction between PRMT2 and AR, Grypari and coworkers sought to determine whether a correlation existed between PRMT2 expression and PCa (175). The authors identified increased PRMT2 cytoplasmic expression in both neoplastic and metastasizing lymph nodes cells compared to healthy prostate tissues (175). The findings in this study were limited to immunohistochemical analysis and patient cohort dataset so the precise roles of PRMT2 in PCa were not elaborated. 1.2.3 Suspected PRMT2 methylation activity The first potential substrate for PRMT2 was identified in 2001 as E1B-AP5, an hnRNP protein that interacts with the adenoviral early protein E1B-55 kDa (176). Kzhyshkowska and coworkers identified a GAR region within the core of the E1B-AP5, that is methylated in vivo by an unknown arginine methyltransferase that co-precipitates with E1B-AP5 (the authors refer to this GAR region as an RGG-box). A recombinant form of the RGG box from E1B-AP5 could be methylated in vitro by human PRMT1 (referred to as HRMT1L2), but not PRMT2 (176). Interestingly, neither enzyme could methylate endogenously expressed E1B-AP5 from cell lysates. The authors performed yeast-two hybrid screening using E1B-AP5 as a bait protein against a HeLa cDNA library and identified the N-terminus (SH3 domain) of PRMT2 as an interaction partner (176). In a separate study, the PRMT2 SH3 domain was identified as an interaction partner with the proline-rich sequences in both SmBâ and Sam68 (hnRNPs) in a protein-domain microarray (30). An interaction between the SH3 domain of PRMT2 and E1B-AP5 was confirmed in vitro by co-IP when anti-E1B-AP5 only co-precipitated full length PRMT2 and not 22 PRMT2ÎSH3 (176). It was also demonstrated by immunostaining that PRMT2 and E1B-AP5 co-localize in the nucleus. Taken together, the authors suggest E1B-AP5 may be a substrate for PRMT2 in vivo but the interaction between the two proteins is likely mediated by a multiprotein complex to stabilize the transient interaction of the SH3 domain (176). Another study in 2009 found that PRMT2 could bind to hnRNP K (an hnRNP associated with nuclear shuttling of DNA), but failed to characterize any methylation activity by PRMT2 (177). These experiments demonstrated a binding interaction between the non-catalytic SH3 domain of PRMT2 and hnRNP proteins but failed to conclusively link PRMT2 to the arginine methylation of hnRNPs. Bona fide PRMT2 catalytic activity was first characterized by the Frankel laboratory in 2009 (35). Using a highly sensitive mass spectrometry technique that can distinguish between and quantify the three mammalian methylarginine species, it was demonstrated that in vitro GST-PRMT2 could deposit MMA and aDMA marks on GST-GAR (glutathione-S-transferase tagged GAR polypeptide derived from fibrillarin (178)) and recombinant histone H4 (25, 35). Compared to PRMT1, PRMT2 had a comparable KM (app) with respect to SAM and histone H4, but the kcat was 800-fold lower than PRMT1 (35). The low methylation activity was corroborated with gel radiography and thin-layer chromatography activity assays. This study also demonstrated that PRMT2 catalytic activity was dependent on the PRMT2 SH3 domain and sensitive to pH and NaCl. It was postulated that the methylation of histone H4 by PRMT2 may be involved in its transcriptional coactivation properties but does not explain its transcriptional repression properties (E2F). Concurrently with the in vitro characterization of PRMT2 methylation activity, the Nabel lab provided evidence that PRMT2 is involved in the arginine methylation of signal transducer and activator of transcription 3 (STAT3) in vivo (179). PRMT1 and PRMT3 methylate Arg-31 of STAT1, a PTM required for the transcriptional activation of STAT1 by promoting DNA-binding, but activity against STAT3 was not characterized (180). The Nabel lab identified that arginine methylation of STAT3 was minimal in cells lacking PRMT2, and significantly elevated in cells transfected with PRMT2, but not PRMT2 lacking the SAM-binding domain (a hybrid PRMT2 made by joining the SH3 domain to the β-23 barrel domain) (179). It is important to note, the PRMT2 SAM-binding domain is in the middle of the protein structure and deletion of this domain will undoubtedly change the entire fold of the protein. The authors provide evidence that PRMT2 enhances STAT3 arginine methylation in vivo, but given the low basal signal observed for STAT3 arginine methylation without PRMT2 expression, there is the possibility that another more active PRMT may be involved. Shortly after the discovery of PRMT2 catalytic activity against histone H4 in vitro, Blythe and coworkers identified an association between PRMT2 and the methylation of histone H3 arginine 8 (181). In this study, Blythe and coworkers were interested in the regulation of the Wnt/β-catenin signaling pathway and how the target genes were regulated during embryogenesis of Xenopus laevis. They identified that β-catenin was responsible for targeting gene promoters for transcription by recruiting chromatin-modifying activity associated with the H3K4me3 and H3R8me2a epigenetic marks (181). When β-catenin was purified from X. laevis embryo lysates, a large macromolecular complex (669 â 2000 kDa) was identified with arginine methyltransferase activity. The authors perform a candidate-based approach that failed to identify an interaction between CARM1, PRMT5, or PRMT6 (canonical H3 binders) and β-catenin, yet GST-PRMT2 could (note: untagged PRMT2 did not copurify with β-catenin) (181). It was further demonstrated that myc-PRMT2 purified from X. laevis was capable of methylating H3R8 in vitro, yet the authors conceded that the activity may be from another copurifying PRMT (181). Regardless of PRMT2 methyltransferase activity, PRMT2 was further shown to associate with target gene promoters in a β-catenin-dependent manner. A unique PRMT2 fusion to the DNA binding domain, ÎNLef1 (Tcf/Lef DNA binding factors that direct β-catenin to gene promoters), was transfected into X. laevis embryos and shown to promote embryonic differentiation (dorsal specification) in the absence of β-catenin (181). When the PRMT2ÎNLef1 fusion was mutated to impair SAM-binding (G145R, G147R identified in: (90)), there was a reduction in the dorsal specification. Taken together, this work demonstrated that PRMT2 associates with gene promoters in vivo and is associated with the H3R8me2a mark. No subsequent study was conducted to determine whether PRMT2 could direct the methylation activity of another PRMT enzyme towards the H3R8 mark. 24 In 2014, Su and coworkers provided some evidence for the in vitro methylation of H3R8 by PRMT2 (14). These authors were investigating a dual Tudor domain containing reader of the H3K4me3/H3R8me2a (or R8me2s) mark, Spindlin1 (SPIN1), using biochemical techniques to characterize the interaction between differentially modified H3 peptides and SPIN1. Using isothermal titration calorimetry (ITC) and differential scanning fluorimetry (DSF), it was demonstrated that SPIN1 associates most tightly to the dually modified H3 peptide bearing K4me3 and R8me2a (with weaker interaction with R8me2s or R8me) (14). While the synthetic peptides used in the binding assays were purchased, the authors also attempted methylation assays using MLL1 (a lysine methyltransferase) against H3 unmodified or R8me2a modified peptides, or using GST-PRMT2 against H3 unmodified or K4me3 modified peptides (14). MLL1 had greater methyltransferase activity against unmodified H3 peptide compared to H3R8me2a, demonstrating crosstalk between the modifications and suggesting the H3K4me3 mark may be deposited first in vivo (14). GST-PRMT2 was shown to methylate H3K4me3 modified peptide to a greater extent than unmodified H3 peptide, but there were several issues with their in vitro methylation assays. First, they did not include a negative control for 3H-SAM with peptide alone for the 8-h incubation at 37 ºC, that must be included since SAM is unstable in vitro and may react with in a non-specific, uncatalyzed manner with nucleophilic residues on the peptides (182). Second, the authors do not include an auto-methylation control of GST-PRMT2 with peptide alone. PRMT enzymes experience auto-methylation, therefore, a control for SAM and enzyme must be included, especially after such a lengthy incubation where auto-methylation may dominate the signal (35, 42). Cavarelli and coworkers demonstrated that PRMT2 has significantly greater auto-methylation activity than it does H3 peptide methylation (reviewed in section 1.2.4) (141). Finally, the authors used GST-PRMT2 bound to resin for their reactions with H3 peptides that were spun down after the reaction and the resin (not the supernatant) was blotted on P8 filter paper. While the GST-resin will be pelleted with the GST-PRMT2, the peptides will not unless bound by the enzyme. Should the assay be repeated, for validation of improved PRMT2 methylation activity against H3K4me3 peptides compared to unmodified H3 peptides, the biotinylated peptides could easily be separated from the enzyme by using avidin resin to separate the 25 H3 peptide for scintillation counting (183). This paper provides biophysical evidence that the SPIN1 reader can bind the H3R8me2a mark, and that crosstalk exists between the H3K4me3 and H3R8me2a marks but fails to provide compelling evidence of the in vitro methylation of H3R8 by PRMT2. The work conducted by Blythe and Su provided some evidence of an interaction between PRMT2 and the H3R8me2a mark that has since been identified as a key driver of an oncogenic phenotype in glioblastoma multiforme (GBM), hepatocellular carcinoma (HCC), and renal cell carcinoma (RCC). PRMT2 mRNA was found to be upregulated in patients with GBM, HCC, and RCC (184â186). Short hairpin RNA (shRNA) targeting PRMT2 was used in primary cell lines for GBM, HCC, and RCC, and shown to inhibit PRMT2 expression (confirmed by Western blot analysis), decrease tumor growth, and inhibit cellular self-renewal properties (184â186). In each study, the groups correlated the expression of PRMT2 with the deposition of the H3R8me2a mark at gene promoters responsible for regulating access to genes associated with the cell cycle, DNA replication, and signaling pathways (184â186). Dong and coworkers were the first to provide some evidence for PRMT2 catalytic activity generating the H3R8me2a mark in GBM cell lines (186). Dong and coworkers generated two PRMT2 SAM-binding mutants, H112Q and M115I (guided by in silico modelling), transfected them in U87 cells and observed that only the H112Q mutant decreased H3R8me2a levels (measured by Western blot analysis) and inhibited cell growth (compared to transfection of wild-type PRMT2). It is important to note that in the absence of functional PRMT2, there was still a low level of H3R8me2a detectable from U87 cells. To validate that PRMT2 was not affecting the activity of PRMT1, the authors mention that PRMT1 inhibition did not impact H3R8me2a levels, but did not evaluate inhibition of other PRMTs (186). Hu and coworkers transfected plasmids encoding the PRMT2H112Q mutant in Huh-7 cells (deficient in PRMT2) or treated Huh-7 cells (overexpressing PRMT2) with MS023 in attempt to inhibit PRMT2 methylation activity in cells (185). Both of these conditions led to decreased tumor volume and reduced the frequency of the H3R8me2a mark (by Western blot analysis), which the authors conclude was attributed to decreased PRMT2 expression or inhibition of PRMT2 catalytic activity (185). MS023 is a Type I PRMT inhibitor that has never been shown to bind to or inhibit PRMT2 activity (131). While the mechanism of 26 PRMT2-mediated regulation of transcription is largely consistent between GBM, HCC, and RCC, the genes transcribed and their effects on tumorigenesis remain unique. These studies suggest that PRMT2 maintenance of the H3R8me2a is the main contributor towards an oncogenic phenotype, but do not address the coactivator/co-repressive properties of PRMT2 towards transcription factors and nuclear receptors (184â186). The authors additionally did not consider whether other PRMTs are involved with the deposition of this mark in association with PRMT2. PRMT2 has also been implicated in the methylation of non-nuclear proteins. In 2018, Hou and coworkers provided biochemical evidence that PRMT2 is involved with the arginine methylation of Cobl, an actin nucleator that regulates neuronal morphogenesis (187). PRMT2 was identified as an interaction partner for Cobl using yeast two-hybrid screening and subsequent coprecipitation analysis that showed the PRMT2 SH3 domain is essential for the interaction between PRMT2 and Cobl (187). The interaction between PRMT2 and Cobl in cells was key for driving the neuronal dendrite formation, which was validated by a total reduction in dendrite formation when shRNA was introduced to inhibit PRMT2 mRNA. A functional PRMT2 SAM binding domain is essential for Cobl arginine methylation since mutation of the SAM binding domain (I153A, L154A, D155A, V156A, G159R, and G161R) inhibited dendrite formation (187). Quantitative coprecipitation and co-IP assays demonstrated that PxxP (x is any amino acid) motifs with flanking positive residues in the Cobl homology domain C-terminus are bound by the PRMT2 SH3 domain (with a KD of 930 nM) (187). Removing the PRMT2 recognition site from Cobl, or mutating the PRMT2 SH3 domain (P92L), resulted in attenuation of Cobl arginine methylation in vivo. Arginine methylation was detected in the extreme C-terminus of the Cobl protein in the WASP homology 2 domain (WH2), responsible for binding actin, directly impacting actin association of Cobl. Using AMI-1 (a pan-PRMT inhibitor) and small interfering RNA (siRNA), the authors demonstrated that arginine methylation of Cobl is reduced but not fully inhibited (187, 188). It is important to note that the Cobl arginine methylation still occurs in the absence of PRMT2, indicating another PRMT may play a role in this interaction. PRMT2 may serve as a recruitment platform that can connect Cobl with another more active PRMT to facilitate arginine methylation. This study further demonstrates the importance of 27 the PRMT2 SH3 domain for protein-protein interactions and provides more evidence implicating PRMT2 with substrate methylation (either directly or indirectly through another PRMT). Another postulated PRMT2 substrate is the human immunodeficiency virus (HIV)-Tat1 protein. The HIV-Tat1 protein is responsible for recruiting the super elongation complex to stalled RNA polymerase II complexes at the promoter sequences of viral-transcripts, to allow expression of viral proteins (155). Inhibiting the association between HIV-Tat1 and the super elongation complex would prevent the HIV virus from escaping the latency phase and keep the virus dormant, thus inhibiting Tat1 association could be a viable HIV therapeutic opportunity. Arginine methylation of HIV-Tat1 by PRMT6 has previously been identified as a PTM that can prevent the association between Tat1 and the super elongation complex (189). In the study conducted by Jin and coworkers, PRMT2 was identified as another PRMT capable of suppressing HIV-Tat1-dependent transcription, using an HIV reporter assay (155). The authors demonstrated in both HEK293 and primary CD4+ cells (healthy or HIV infected) that PRMT2 overexpression promoted HIV latency, but a PRMT2 catalytic mutant (mutation of the glutamic acids composing the double E loop to glutamine) did not. Co-IP and GST-pulldown assays revealed the PRMT2 SH3 domain was key for the interaction with Tat1 (155). Additionally, the authors purified flag-tagged PRMT2 from HEK293 and demonstrated in vitro activity of the complex against Tat1 at Arg-52 and Arg-53, like PRMT6. In fact, it was demonstrated that the presence of both PRMT2 and PRMT6 resulted in a synergistic increase in Tat1 arginine methylation (155). 1.2.4 PRMT2 structure In 2017, the first crystal structure of PRMT2 was solved (141). Cavarelli and coworkers were able to generate the crystal structures for Danio rerio PRMT2 (zPRMT2 â D. rerio is commonly referred to as zebrafish) in complex with either sinefungin or SAH, as well as Mus musculus PRMT2 (mPRMT2) in complex with either SAH or Cp1 (a synthetic adenosine scaffold connected to a guanidine). zPRMT2 and mPRMT2 orthologs share 69.2% and 85.3% sequence similarity with human PRMT2, respectively, with the greatest difference in sequence between the orthologâs SH3 and N-terminal domains. The structures demonstrate high similarity in their structural fold with a root mean square deviation (RMSD) 28 of 0.68 Ã between zPRMT2 and mPRMT2 (141). A structural overlay of mPRMT2 with PRMT1 and PRMT6 reveals a highly conserved structural fold, with the greatest observed differences between the mobile N-termini and dimerization arms (Figure 1.5, Table 1.1). Like the other PRMT enzymes, PRMT2 also forms homodimers. The PRMT2 homodimer is mediated through contact of a dimerization arm of one subunit of a homodimer making reciprocal contacts with the other subunit to form an overall toroidal shape where the two active sites face inward and across from one another (141). Homodimers were confirmed as the major oligomeric species for mPRMT2 through small angle X-ray scattering (SAXS) and crystallographic evidence (141). Figure 1.5. Pairwise structural alignment of PRMT1, PRMT2, and PRMT6. Structure Pairwise Alignment (PDB) between PRMT1 (PDB ID: 1OR8), mPRMT2 (PDB ID: 5FWA), and PRMT6 (PDB ID: 5WCF) rendered using BIOVIA Discovery Studio 2021. PRMT2 (PDB: 5FWA)PRMT1 (PDB: 1OR8)PRMT6 (PDB: 5WCF)Cofactor-binding domain/Rossman foldBeta-barrel domainDimerization armN-terminusDouble E LoopC-terminus29 Table 1.1. Measures describing the structural alignment of PRMT1 and PRMT6 against PRMT2 using pairwise structural alignment. Visualized alignment of macromolecular structures shown in Figure 1.5. TM-score represents topological modelling score. Protein Entry Chain RMSD TM-score Identity Equivalent Residues Sequence Length Modelled Residues PRMT2 5FWA A - - - - 446 339 PRMT1 1OR8 A 1.85 0.87 32% 312 340 313 PRMT6 5WCF A 2.33 0.85 37% 313 376 319 Unfortunately, the crystal structure for both mPRMT2 and zPRMT2 did not include electron density for the SH3 domain, the authors attribute this to a lack of structural rigidity (141). A structure of the PRMT2 SH3 domain (without the methyltransferase and β-barrel domains) was solved by nuclear magnetic resonance spectroscopy (NMR), revealing a conserved fold of two perpendicular β-sheets formed by five antiparallel β-strands, shared amongst other SH3 domains (PBD: 1X2P â not published) (190). The PRMT2 SH3 domain is a ~60 amino acid protein-binding module that is conserved amongst signal transduction proteins (191). There are different motifs that can be recognized by distinct SH3 domains, such as Class I ligands bearing an RxxPxxP motif (where x is any amino acid) or Class II ligands bearing the xPxxPxR motif, but the specificity of the PRMT2 SH3 domain is unknown (192). Other studies have identified non-canonical motifs that can be bound by the SH3 domain, that contain cationic arginine and lysine residues in sites normally occupied by proline, as demonstrated by X-ray crystallography (193). The AlphaFold structure of PRMT2 shows the SH3 domain connected to the catalytic methyltransferase domain by a flexible linker (Figure 1.6) (194). There is low confidence in the conformation of the flexible linker, indicating the exact positioning of the SH3 domain is unknown. 30 Figure 1.6. Predicted structure of full-length PRMT2. Model of the full-length human PRMT2 (1-433) structure with colors showing model confidence by AlphaFold. Images rendered by Biovia Discovery Studio 2021. A31 Within the PRMT2 active site, there are several conserved motifs with respect to other PRMTs, like the double-E loop and the THW loop in proximity to the guanidino constituent of Cp1, and the positioning of motif I within the Rossmann-like fold âILDVxxxGxGâ with respect to the SAM binding pocket (Figure 1.7A). The IDVxxxGx motif (motif I) has been the target of site-directed mutagenesis to impair SAM binding in a few studies previously discussed (90, 165, 187). A closer examination of the SAM binding site reveals that motif I does not form non-covalent contacts with SAH directly, but instead appears to serve as a structural anchor to hold the SAM binding site together (Figure 1.7A). Mutagenesis of motif I to alanine residues will likely impair SAM binding by disrupting the intramolecular fold of the SAM binding site, potentially leading to greater protein instability. Other mutations, like H112Q or M115I (with respect to human PRMT2 sequence), mutate direct contacts with SAH and minimally impact the steric and hydrogen bonding of the PRMT2 active site (Figure 1.7A) (186, 195). Pak and coworkers demonstrated that mutating one of the two glutamic acids composing the double E loop (E220Q with respect to human PRMT2 sequence) abolished PRMT2 catalytic activity and the crystal structure of mPRMT2 confirms this residue contacts the guanidino group of Cp1 (Figure 1.7B) (19, 141). 32 Figure 1.7. mPRMT2 binding interactions with SAH and Cp1. (A) Zoomed in view of the Rossmann-like fold bound to SAH (carbon atoms in purple) highlighting residues composing motif I (153-ILDVxxGxG-159) and residues H124 and M127 (carbon atoms in yellow) (PDB ID: 5FUL). Dashed lines represent 33 non-covalent interactions with SAH, H124 forms a C-H bond to 3âOH of SAH, M115 forms a Sulfur-X bond to the sulfur atom in SAH (B) Zoomed in view of the Rossmann-like fold bound to Cp1 (carbon atoms in purple) highlighting the glutamic acids composing the double-E-loop (E223, E232) and the THW loop (380-THW-382) (PDB ID: 5FWA). Dashed lines represent non-covalent interactions with Cp1, E223 and E232 both form conventional hydrogen bonds with the terminal and internal nitrogen atoms of the guanidino ring. Images rendered using BIOVIA Discovery Suite 2021. In addition to solving the crystal structure of non-human PRMT2 orthologs, Cavarelli and coworkers also characterized the activity and binding phenomenon of mPRMT2. Cavarelli and coworkers corroborated previous findings that a PRMT2 truncation lacking the SH3 domain had reduced catalytic activity (141). Like the in vitro characterization of human PRMT2 catalytic activity, Cavarelli and coworkers also relied on gel radiography to evaluate mPRMT2 catalytic activity against histone H3 and H4 peptides, PRMT2 (auto-methylation), and repressor splicing factor 1 (RSF1). RSF1 was identified as a PRMT2 co-purifying protein from the insect expression cell line, Spodoptera frugidepa 9, that could be methylated by PRMT2 (141). Michaelis-Menten kinetics were not performed on mPRMT2, so a comparison of the catalytic activities of mPRMT2 and human PRMT2 could not be conducted. It was demonstrated that mPRMT2 displayed greatest activity towards RSF1, followed by itself (auto-methylation), and had very low methylation activity against the histone peptides (141). While RSF1 does not exist in human cells, the authors predict that the substrate PRMT2 has greatest activity against could be similar to RSF1 (141). Using RSF1 as a substrate, Cavarelli and coworkers showed that both SAH and Cp1 could inhibit PRMT2 catalytic activity with an IC50 of 18.3 and 16.3 µM, respectively. These results were corroborated with a thermal shift assay of PRMT2 in the presence or absence of either SAH of Cp1, demonstrating the molecules (at 1 mM) significantly thermally stabilized PRMT2 (141). The thermal shift technique has been used as a tool to evaluate how PRMTs interact with potential ligands (24, 131, 196â199). This study marks a significant advancement for understanding PRMT2 biochemistry as it provides the structural blueprints for PRMT2 active site architecture (conserved amongst PRMT2 orthologs), while employing novel in vitro techniques to characterize mPRMT2. The structure of human PRMT2 has not been solved, but the work conducted by Cavarelli and coworkers provided the foundations for the work completed in chapters 3 and 4 of this thesis. 34 1.2.5 PRMT2 proteomics investigations While studies described above highlight individual PRMT2 interactions, proteomic investigations have identified a plethora of additional PRMT2 interactions. In 2017, Vhuiyan and coworkers published a study on the interaction between the PRMT2 SH3 domain and splicing factors, including Src associated in mitosis 68 (Sam68) protein, a regulator of the alternative splicing for BCL-X (200). First, the authors performed a GST-pulldown experiment using GST-SH3 (from PRMT2) as bait and HeLa cell lysate as prey, in addition to running GST only and GST-SH3 (Abl) as negative controls. Upon comparing the proteins pulled down using the PRMT2 SH3 domain to the negative controls (by mass spectrometry proteomics identification), there was a significant enrichment of more than 30 splicing-related proteins, including hnRNP U (E1B-AP5 analogue) and hnRNP K, validating previous interactions (176, 177, 200). The authors further characterized the interaction between PRMT2 and Sam68, a PRMT1 substrate involved with the alternative splicing of BCL-X (65, 201). An SH3-dependent interaction between PRMT2 and Sam68 was confirmed through reciprocal co-IP experiments (200). Using immunofluorescence on transfected HeLa cells (with mCitrine PRMT2 or mCitrine PRMT2ÎSH3), it was further demonstrated that PRMT2 and Sam68 colocalize primarily in the nucleus. The HeLa cells were then treated with AdOx (adenosine dialdehyde), a molecule that inhibits SAH hydrolase (SAHH) that results in a cellular accumulation of SAH and increased inhibition of methyltransferase enzymes, which disrupted PRMT2 nuclear colocalization with Sam68 and resulted in cytoplasmic localization only (200). In the absence of the SH3 domain, PRMT2 and Sam68 colocalized independent of AdOx treatment. Vhuiyan and coworkers also established that PRMT2 increased the BCL-X(L)/BCL-X(s) ratio in cells stimulated with LPS of TNF-α (200). BCL-X(L) is the anti-apoptotic splice variant of BCL-X and BCL-X(s) is the apoptotic splice variant, therefore the increase in ratio signals apoptosis resistant, consistent with the PRMT2 effect on NF-κB signaling pathway (165). There has been a significant rise in the number of PRMT2 interaction partners through proteomic investigations of interaction networks in different cell lines (149, 202, 203). These mass spectrometry approaches have involved tagging libraries of human proteins with hemagglutinin (HA) or Flag affinity 35 tags, transfecting them into the cell line of interest, then performing affinity pulldown on the bait proteins and using proteomics to identify interacting partners. PRMT2 was one of the bait proteins used for pull down experiments and multiple proteins could co-purify with PRMT2, indicating protein-protein interactions (149, 202, 203). Some of these PRMT2 co-purifying proteins were also used as bait proteins and were able to pulldown endogenous PRMT2, demonstrating reciprocity in the protein-protein interaction. Amongst the interactions, post-translational modification writers, SH3 interactors, splicing factors, and transcription factors were identified (149, 202, 203). Graphical representations of the protein-protein interaction networks associated with PRMT2 can be found in Figure A2-3. The STRING PRMT2 protein-protein interaction network (Figure A3) also included co-expression as an interaction node where PRMT2 gene expression was linked to dysregulations in L-arginine metabolism in addition to learning disorders (associated with chromosomal deletion or duplication of chromosome 21) (204â210). 1.2.6 PRMT2 heteromeric interactions Bona fide PRMT2 methylation activity has been difficult to prove conclusively since the inhibition of cellular PRMT2 does not fully attenuate suspected methylation activity (e.g., on H3R8, E1B-AP5, Cobl) (176, 181, 185, 186). It has been postulated that PRMT2 can interact with other PRMTs and promote their catalytic activity. The first evidence demonstrating PRMT2 interactions with other PRMTs came in 2009 when Hermann and coworkers conducted an in vivo characterization of all mammalian PRMT enzymes (39). To characterize the cellular localization of the PRMT enzymes, they transfected GFP-tagged variants of all PRMTs (separately) into HEK293 cells and observed the fluorescent signal (39). PRMT2 was shown to localize primarily in the nucleus, consistent with other studies (160, 161, 164, 200). GFP-PRMT2 overexpressing HEK293 cells were transfected with the other mammalian PRMTs, lysed, and then PRMT2 was immunoprecipitated to determine whether it could pulldown the other PRMTs (39). PRMT2 was able to pulldown PRMT8 (and PRMT1 and PRMT6 to a lesser extent) (39). The PRMT2-PRMT8 interaction was mediated through the PRMT2 SH3 domain and a proline-rich domain within the N-terminus of PRMT8 (211). The study by Hermann and coworkers also demonstrated that the same PRMT2 immunopurified from HEK293 had weak enzymatic activity in comparison to 36 PRMT1, but did not explore whether an additional PRMT copurified with PRMT2 that was responsible for the methyltransferase activity (39). To address whether PRMT2 could interact with other PRMT enzymes, the Frankel lab conducted a study on the heteromeric interactions between PRMT1 and PRMT2 (19). A heteromeric interaction between PRMT1 and PRMT2 was validated by reciprocal co-IP experiments, both in vitro and in HeLa cells overexpressing PRMT1 and PRMT2 (19). The authors then transfected HeLa cells with the N-terminus of mCitrine (mCitN) fused to PRMT1 and the C-terminus of mCitrine (mCitC) fused to PRMT2, that resulted in reconstituted citrine fluorescence in cells, demonstrating a protein-protein interaction using biomolecular fluorescence complementation (BiFC) (19). It was further demonstrated that PRMT2 could augment the catalytic activity of PRMT1 towards histone H4, in vitro, and overexpression of PRMT1 and PRMT2 in HeLa cells resulted in synergistic augmentation of total cellular arginine methylation (19). Since all types of methylarginine were augmented, including sDMA, this indicates that PRMT1 and PRMT2 overexpression impacted either PRMT5 or PRMT9 activity in HeLa cells. The authors demonstrated that the interaction between PRMT1 and PRMT2 did not depend on the catalytic activity of PRMT2, since PRMT2E220Q could co-IP PRMT1 and augment its catalytic activity to the same extent as wild-type PRMT2 (PRMT2E220Q was validated as a PRMT2 catalytically inactive mutant through ultra-high performance liquid chromatography with tandem mass spectrometry (UPLC-MS/MS analysis)) (19). To identify structural determinants for the interaction between PRMT1 and PRMT2, the authors performed BiFC and co-IP of HeLa cells containing PRMT1 or PRMT2 constructs lacking either the dimerization arms, the PRMT2 SH3 domain, or with catalytically inactivating mutations (PRMT1E153Q, PRMT2E220Q) (19). Co-IP demonstrated that deletion of the PRMT2 dimer arm impaired the interaction between PRMT1 and PRMT2, but BiFC demonstrated weak fluorescence reconstitution, leading to inconclusive results. The N-terminus of mCitrine fused to a PRMT2 truncation lacking the SH3 domain (mCitN-PRMT2ÎSH3) was capable of reconstituting fluorescence with mCitC-PRMT1, indicating the SH3 domain was not responsible for the interaction between PRMT1 and PRMT2. PRMT1 catalytic activity was required for BiFC signal with PRMT2 since neither PRMT1E153Q, nor 37 PRMT2, in AdOx-treated cells, could reconstitute citrine fluorescence (19). These data provide strong evidence that PRMT2 can physically interact with PRMT1 in cells and in vitro to modulate its activity. PRMT2 was recently shown to be involved in the formation of a ternary complex with CARM1 and bromodomain containing protein 4 (BRD4) (154). In exploring the molecular mechanisms connecting BRD4 to transcriptional control, Liu and coworkers investigated whether PRMT enzymes could interact with and methylate BRD4. Reciprocal IP experiments demonstrated that BRD4 could interact with both PRMT2 and CARM1 (154). CARM1, and to a lesser extent PRMT2, when overexpressed in MCF7 cells, increased the level of MMA formation on BRD4, yet in vitro, only CARM1 could methylate BRD4 (using 3H-SAM based in vitro methylation assay) (154). Through site-directed mutagenesis, it was identified that three non-consecutive arginine residues (3R) between the bromodomains of BRD4 were the target of CARM1 methylation in cells. The 3R site of BRD4 does not conform to the PGM motif preferred by, indicating that PRMT2 may modulate the substrate specificity of CARM1 (154). It was further demonstrated that arginine methylation of BRD4 is required for BRD4 to bind acetylated histones and recruit transcriptional machinery at active promoters and enhancers to promote transcription. The inhibition of BRD4 arginine methylation, through shRNA silencing of PRMT2/4, knock-in mutants of BRD4 3R to K (3RK), or treatment of MDA-MB-231 cells with TP-064 (a CARM1-selective inhibitor) dramatically reduced expression of genes regulating DDR pathways (154). It was further demonstrated through MDA-MB-231 cells (expressing BRD4WT, BRD43RK, or treated with shPRMT2/4 shRNA) and MDA-MB-231 xenograft mice models that inhibition of BRD4 arginine methylation significantly decreased cellular proliferation and tumor growth (154). This study clearly demonstrates that PRMT2 can augment the catalytic activity of CARM1 towards BRD4. In addition to the evidence of PRMT2 augmenting PRMT1 and CARM1 activity, it was also demonstrated that PRMT2 can synergistically increase arginine methylation of the HIV-Tat1 protein with PRMT6 (reviewed in 1.2.3) (155). It is unknown how PRMT2 interacts with these proteins and further characterization is required. 38 1.3 Hypothesis and research objectives PRMT2 has been implicated in a plethora of biological functions through protein-protein interactions and its postulated methylation activity, yet in vitro validation of these interactions is lacking. A major barrier with analyzing the low catalytic activity of PRMT2 or characterizing its biophysical interactions is cost. Screening the activity of low activity methyltransferase enzymes requires sensitive equipment to detect a signal that are expensive and often requiring multiple days of sample preparation. Characterizing protein-protein and protein-ligand interactions requires expensive instrumentation that can measure thermodynamics or binding phenomenon of a system. The first research goal of this dissertation was to develop in vitro cost-effective techniques to characterize the biochemical function of PRMT2. The second major goal of this dissertation was to investigate the mechanisms of how PRMT2 facilitates methylation of histone proteins in concert with other PRMTs. My formal research hypothesis is as stated: PRMT2 can interact with and alter the substrate specificity of PRMT1 to promote the formation of specific epigenetic marks. 39 Chapter 2. Protein arginine N-methyltransferase activity determination with filter binding and phosphor screening (FBAPS) assay 1 1 A version of this section has been published. Rowley, M. J., Taylor, S. A., and Frankel, A. (2022) Protein arginine N-methyltransferase activity determination with filter binding and phosphor screening (FBAPS) assay. Anal. Biochem. 653, 114778 I designed and conducted all experiments, parameterization, data analysis, and protein purification. Shona A. Taylor performed preliminary experiments, which does not appear in this thesis, and contributed helpful points of discussion. Dr. Adam Frankel and I designed experiments, analyzed data, and wrote and edited the manuscripts. The PRMT2 related content was not included in the published manuscript. 2.1 Introduction Human protein arginine-N-methyltransferases (PRMTs) catalyze the transfer of a methyl group from S-adenosyl-L-methionine (SAM) to terminal guanidino nitrogen atoms on arginine-containing polypeptides, forming methylarginine and S-adenosyl-L-homocysteine (SAH) (8). These enzymes regulate various cellular signalling pathways and have been identified as prominent therapeutic targets against several cancers including brain, breast, and hepatic cancers (3, 186). The inhibition of PRMTs in cancer models through siRNA knockdown or small molecules has been shown to halt tumor growth and increase survival (131, 186, 212). A major bottleneck in developing therapeutics against PRMT enzymes is the limited throughput in traditional screening technologies designed for measuring methyl group transfer. Studying the kinetics of methyl group transfers can be done through one of several techniques: 1) tracking the transfer of radiolabelled methyl groups, 2) using mass spectrometry assays that can detect either intact whole protein methylation or acid-hydrolyzed products of methylated polypeptides, or 3) measuring SAH or SAH metabolite production (35, 213â215). The usage of radiolabelled SAM has dominated the PRMT field due to its high sensitivity and ease of use. PRMT kinetics assays such as the P81 filter binding assay (22) and gel radiography (35) both rely on the use of radiolabelled SAM cofactor (14C-methyl or 3H-methyl) as the source of the transferable, 40 isotopically-labelled methyl group that becomes enzymatically appended to arginine residues in polypeptide substrates. Substrate-bound radioactivity can be detected by scintillation counting or phosphor imaging after separation from unspent cofactor. The P81 filter binding assay traditionally involves blotting reactions with radiolabelled cofactors (14C-SAM for methyltransferases; ð¾-32P-ATP for kinases) onto 1.0-cm2 pieces of paper, followed by washing away unreacted SAM prior to liquid scintillation counting (216). Advances in P81 filter binding have included increasing throughput via a slot blot apparatus, allowing for 96 reactions to be blotted simultaneously (214). Gel radiography involves separating methyltransferase reactions electrophoretically on an SDS-PAGE gel, which is stained to visualize polypeptides, dried, and subsequently exposed to a storage phosphor screen for phosphor imaging (217). Both techniques are limited in throughput and not feasible for large-scale screening efforts. Recent progress in the field has led to the development of high-throughput PRMT assays using radiolabelled cofactor. The scintillation proximity assay (SPA) employs specialized scintillation beads that emit light upon excitation by radioactive decay without the need to separate unreacted SAM from methylated substrate (218), and has yielded impactful results in identifying PRMT chemical probes (219, 220). The ZipTip® assay uses specialized pipette tips containing C18 resin used to separate peptide substrates from unreacted SAM; processing of reactions with a multi-channel pipette can increase throughput (213). Given the additional costs associated with SPA kits and ZipTips, we sought to develop a more economical assay. This work aims to improve the throughput of the traditional P81 filter binding assay while eliminating the need to use expensive and environmentally unfriendly scintillation fluid by using phosphor imaging instead. The filter binding and phosphor screening (FBAPS) assay yields reliable data while eliminating added expense. A similar method has been described for protein kinases, using P81 filter binding and phosphor screening with ð¾-32P-ATP as the radiolabel (221). Protein arginine N-methyltransferase 1 (PRMT1) and coactivator-associated arginine methyltransferase 1 (CARM1) were selected to develop the FBAPS assay given their well-characterized activities and established inhibitors (SAH, MS023, Diamine 12, EPZ020411) (24, 47, 131). To determine the accuracy of this assay, two kinetic characterizations were performed: KM determinations for PRMT1 41 with glutathione-S-transferase fused to the glycine- and arginine-rich polypeptide (GST-GAR) and CARM1 with GST fused to poly(A) binding protein 1 (437-488) (GST-PABP1(437-488)) (222); IC50 and estimated KI determinations were made for SAH, Diamine 12, MS023, and EPZ020411 against both enzymes. We additionally evaluated the applicability of FBAPS towards investigating a low-activity protein arginine methyltransferase by measuring the linear range of PRMT2 methylation of repressor splicing factor 1 (RSF1), thereby identifying important limitations with this assay. 2.2 Methods 2.2.1 Reagents SAH was purchased from MilliporeSigma (A9383) and dissolved in 0.5 mM HCl (to prevent hydrolysis) to make a 2.0-mM stock. MS023 was received from Chemical Probes (Chemical Probes Portal ID: CPP36) (www.chemicalprobes.org) and dissolved in DMSO to make a 1.0-mM stock. EPZ020411 was purchased from Cayman Chemicals (Item No. 19160), which was dissolved in water to make a 11.3-mM stock. Diamine 12, a gift from Dr. Eric Therrein, was dissolved in DMSO to make a 20-mM stock. The radiolabelled cofactor S-adenosyl-L-[methyl-14C]-methionine (58 mCi mmolâ1) was purchased from PerkinElmer (NEC363050UC). 2.2.2 Protein expression and purification Cloning and expression of PRMT1, PRMT2, and CARM1 in pET28a(+) (Novagene), GST fusions of GAR and PABP1(437-488) in pGEX-2T (Cytiva), and RSF1 in pET15b (Novagene) have been described previously (147, 223, 224). CARM1, PRMT1, PABP1, and RSF1 were transformed and expressed in Escherichia coli BL21(DE3) pLysS (Stratagene). GST-GAR was transformed and expressed in Escherichia coli DH5α (Stratagene). PRMT2 was transformed and expressed in Escherichia coli Arctic Express (DE3) (Stratagene). In brief, cells were grown in LB media to an OD600 of 0.6 and protein expression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) at 16 ºC for 16 h. Cells were harvested via centrifugation (10000 x g, 4 ºC, 15 min) and cell pellets were frozen at -80ºC. Cell pellets containing hexa-histidine tagged PRMT were thawed on ice in a lysis buffer [2 mL/g wet weight of cells; 50 mM HEPES-KOH, 1 M NH4Cl, 10 mM MgCl2, 10 mM imidazole, 0.1% Triton X-100, 0.1% 42 lysozyme, 25 U mL-1 DNAse I, 1.0 mM phenylmethanesulphonyl fluoride (PMSF), 7 mM β-mercaptoethanol (BME), and 1.0 mM EDTA-free protease inhibitor cocktail (PIC), equilibrated to a final pH of 7.4] and incubated for 1 h at 25 ºC before further lysis via sonication (ten 30 s pulses at 50% duty cycle with 30 s pauses, using a Branson Sonifier 450 at 4 ºC). Soluble proteins were separated by centrifugation (35000 x g, 4 ºC, 1 h) and filtered through a 0.22 µM low protein binding PVDF membrane (Millex). Clarified lysates containing hexa-histidine tags were applied to pre-equilibrated Ni-NTA resin or HisTrap FF column (GE healthcare) in wash buffer [50 mM HEPES-KOH, 1 M NH4Cl, 10 mM MgCl2, 10 mM BME, 10 mM imidazole, 1 mM PMSF, equilibrated to a final pH of 7.6] and eluted using stepwise imidazole elution buffers or over a gradient of 50 â 400 mM imidazole when performing column elution (elution buffer had same composition as wash buffer except for 100-400 mM imidazole). Fractions containing the protein (confirmed by 10% SDS-PAGE) were applied to a pre-equilibrated HiLoad26/600 Superdex 200 pg size-exclusion column (SEC) in wash buffer [50 mM Tris-HCl, 100 mM NaCl, 1 mM DTT, pH 8.0]. Column purification performed using AKTA FPLC basic and protein elution monitored by A280 UV-Vis trace. Eluted fractions containing the desired protein (evaluated by 10% SDS-PAGE analysis) were pooled and concentrated in a storage buffer (100 mM HEPES-KOH pH 8.0, 200 mM NaCl, 1 mM DTT, 10% glycerol, 2 mM EDTA) using Amicon Ultra 15 mL filters [30-kDa molecular weight cut-off (Millipore)]. Cell pellets containing GST-tagged proteins (GAR and PABP1) were lysed using the same protocol as listed above except PBS lysis buffer [2 mL/g pellet; 1x PBS, 5% glycerol; 1 mM EDTA; 1 mM EGTA, 0.1% Triton X-100, 0.1% lysozyme, 25 U mL-1 DNAse I, 1.0 mM phenylmethanesulphonyl fluoride (PMSF), 7 mM β-mercaptoethanol (BME), and 1.0 mM EDTA-free protease inhibitor cocktail (PIC), equilibrated to a final pH of 7.3] was used. Following separation of soluble proteins, GST-tagged proteins were injected to a pre-equilibrated 5 mL GST-Trap FF column (GE healthcare) (pre-equilibrated in PBS pH 7.3 and 1 mM DTT) and eluted with reduced glutathione (50 mM Tris-HCl, 10 mM reduced glutathione, 1 mM DTT, pH 8). 43 All protein concentrations were quantified by the Bradford assay (Quick Startâ¢ Bradford Protein Assay; BioRad) (225). 2.2.3 Enzyme assay Unless stated otherwise, enzyme assays were performed with 1.0 µM PRMT1, 11.3 µM GST-GAR, and 50 µM 14C-SAM (3.82 DPM µmol-1-) for 60 min, or 1.0 µM CARM1, 10 µM PABP1(437-488), and 50 µM 14C-SAM for 120 min in methylation buffer (50 mM HEPES, pH 8.0, 10 mM NaCl, 1.0 mM DTT) at 37 ËC (20-µL final reaction volume). PRMT2 linear range experiments were performed with 1.0 µM PRMT2, 15 µM RSF1, and 50 µM 14C-SAM in methylation buffer at 37 ËC for -10 h. To terminate reactions, they were spotted in quadruplicate (2.0 µL spotted twice for a total of 4 µL) under vacuum onto Whatman P81 phosphocellulose filter paper (Fisher Scientific 3698â915) clamped into a BioRad Bio-BlotTM Apparatus with a piece of Whatmann 3 MM blotting paper underneath. Methods for radioactivity measurement are described below. The conditions used are within the linear range of the enzymes (Figure A4). Linear ranges determined by incubating PRMT1 or CARM1 reactions from 0-360 min with 11.5 µM GST-GAR or 20 µM GST-PABP1(437-488), respectively. The time-dependent rates were analyzed with simple linear regression from which the apparent linear ranges were estimated using GraphPad Prism 8.4.3. 2.2.4 Scintillation counting For data analyzed by scintillation counting, the P81 filter paper was washed again with absolute methanol (10-20 s), allowed to air dry, and then stained with 200 mL 0.5% Coomassie Brilliant Blue G-250 in 7% acetic acid (v/v; 0.22-µm filtered) for 30 min at ambient temperature with gentle shaking. Excess stain was rinsed off with water and the filter paper was destained in 200 mL of 7% acetic acid (v/v) for 30 min. The visible spots indicating protein were excised and submerged in 2 mL Scintiverseâ¢ BD Cocktail (Fisher Scientific, SX18-4). Disintegrations per minute (DPM) were detected for 1 min using a Beckman LS6500 liquid scintillation counter. 44 2.2.5 Densitometry and data analysis Washed and dried filter papers were exposed to a storage phosphor screen for either 1 or 2 h (PRMT1 or CARM1 reactions, respectively). Radiographic images were generated on storage phosphor plates using a Sapphire Biomolecular Imager set at 200-µm resolution, highest scan speed, and an intensity setting of 3. Densitometry analysis was performed by Image Studio Lite Ver 5.2. Images were brightness- and contrast-adjusted to avoid oversaturation. Each spot was analyzed using the Analysis feature on Image Studio Lite where the area was kept constant for each spot and background (vide infra) was subtracted using the median function. After squares were placed over approximate dot locations, the auto-adjust feature was applied to find the exact pixel density (minimum finding threshold set to lowest intensity and search area extension set to four notches from the minimum). Background signal from enzyme-only control reactions (enzyme and SAM only) was subtracted from data points to normalize data sets. 2.2.6 SAM standards and unit conversions Quantitation of a storage phosphor image from an FBAPS assay requires a set of radioactive standards that encompass the activity range of the methyltransferase assayed. Two series of 14C-SAM dilutions were spotted onto strips of P81 filter paper: one capturing 250-20000 DPM for PRMT1 activity calibration, and the other capturing 250-5000 DPM for CARM1 activity calibration. Each strip was placed on the top left side of the P81 filter paper during phosphor imaging as a reference. 2.2.7 KM and kcat determination Methylation reactions for PRMT1 (200 nM) towards 0-11.5 µM GST-GAR and for CARM1 (200 nM) towards 0-50 µM GST-PABP1(437-488) were used to generate rate curves for KM determination. The observed reaction rates were fitted to the Michaelis-Menten equation (1) from which the apparent KM and Vmax were estimated using GraphPad Prism 8.4.3. When substrate inhibition was suspected, the observed reaction rates were fitted to the substrate inhibition equation (2). The apparent kcat values were calculated by converting the phosphor signal using radioactive standards described in 2.6 to DPM, which were converted to µCi (1 µCi = 2.22 x 106 DPM) so that µmol s-1 for each enzyme could be calculated 45 based on the specific activity of 14C-SAM; these values were then divided by total enzyme concentration. Statistically significant differences were those that yielded a p-value < 0.05 in a paired t-test with Welchâs correction. (1) ð£ = ðððð¡[ð¸]ð[ð]ð¾ð+[ð] (2) ð£ = ðððð¡[ð¸]ð[ð]ð¾ð+[ð](1+[ð]ð¾ð¼) 2.2.8 IC50 determination The inhibitory effects of SAH, MS023, and EPZ020411 against PRMT1, and SAH, MS023, and Diamine 12 against CARM1 were determined to validate the performance of the FBAPS assay. For each inhibitor tested, a vehicle-only control was performed (where the enzyme reaction was incubated with the solvent used to solubilize the highest concentration of inhibitor tested â generally 1% DMSO). In all instances, the vehicle-only control did not significantly inhibit catalytic output of the PRMT reactions. Inhibition experiments were performed in triplicate, on three separate days (n = 9). The resulting data were normalized by converting raw phosphor signal to normalized phosphor signal using the following formula: (3) ð¦ = (ððð¥ððð¤+ðððððð¤ ððð¥ððð+ðððððð) (ð¥ â ððð¥ððð) + ððð¥ððð¤ The observed activity rates were fit to the four-parameter logistic equation (4) from which the half-maximal inhibitory concentration (IC50) was derived: (4) ð¦ = ðµðð¡ð¡ðð+(ðððâðµðð¡ð¡ðð)(1+[ð¼ð¶50]ð¥)ð»ððð ððððð where x is inhibitor concentration, and y is phosphor signal. Curve fitting was performed using GraphPad Prism 8.4.3. The apparent KI values were estimated using equation (5) (24). Statistically significant differences were those that yielded a p-value < 0.05 in a paired t-test with Welchâs correction. (5) ð¾ð¼ = ð¼ð¶501+[ð]ð¾ð 46 2.3 Results and discussion 2.3.1 FBAPS assay conditions Methylation reactions were spotted on dry P81 filter paper clamped in a dot blot apparatus according to a loading key (Figure 2.1A), and then removed and washed with 100 mL NaHCO3/Na2CO3 pH 9.0 buffer until scintillation counts of wash buffer fell below 100 DPM (five 100-mL washes; Figure A5) prior to drying for 30 min at 37 ËC. Proteins were visualized on the P81 filter paper by Coomassie blue staining (Figure 2.1B) (226). β-emission from 14C-methylated arginine proteins captured on the P81 filter paper were analyzed by storage phosphor screening (Figure 2.1C) and densitometry analysis (Figure 2.1D). We were able to quantify PRMT activity by converting phosphor signal to DPM with the aid of 14C-SAM serial dilutions of known specific activity (Figure 2.2). Absolute quantification of PRMT activity required that the 14C-SAM quantification standards encompassed the entire activity range for the enzyme used: 2500-20000 DPM or 250-5000 DPM for PRMT1 or CARM1, respectively. This activity range must be determined by scintillation counting prior to FBAPS analysis if absolute quantification is required (e.g., kcat determination). For rapid KM and IC50 determination, absolute quantification and the use of SAM standards are not essential. 47 Figure 2.1. FBAPS assay layout and output. (A) Loading key for spotting methylation reactions (n = 4) into 96-well dot-blot apparatus containing controls in row A: radiolabelled SAM alone, with substrate, or with enzyme; rows B-D: reactions with increasing substrate; rows E-H: reactions with increasing inhibitors. (B) Coomassie blue-stained P81 filter paper blotted with reactions in (A). (C) Phosphor image after a 2-h exposure of P81 filter paper in (B). (D) Heat map showing the densitometry values obtained from the phosphor image in (C). 48 Figure 2.2. Quantitation of phosphor signal. Phosphor images from two sets of 14C-SAM dilutions (top): (A) 23.5-295 pmol 14C-SAM (1-h exposure), and (B) 1.29-48.3 pmol 14C-SAM (3-h exposure) are shown with their corresponding phosphor signal and DPM from the isotopic label (bottom). Linear regressions and corresponding correlation coefficients (R2) are shown. 2.3.2 Enzyme kinetic determinations using the FBAPS assay To evaluate the effectiveness of the assay, we compared KM values using data originating from phosphor signals and scintillation counts (Figure 2.3). The KM values obtained for PRMT1 with GST-GAR (Table 2.1) showed no statistically significant difference (p = 0.1421) between the two methods. Similarly, there was no statistically significant difference between the kcat values determined for PRMT1 between both methods (p = 0.1525 for PRMT1). Evaluation of the kinetic parameters for CARM1 with PABP1(437-488) were complicated since substrate inhibition was observed at higher peptide concentrations (Figure 2.3B). Substrate inhibition was confirmed by fitting the observed reaction rates to both substrate inhibition (R2=0.8582) and Michaelis-Menten equations (R2=0.500). Substrate inhibition is common when performing nanomolar PRMT reactions and can be overcome by increasing concentration of enzyme or avoiding inhibitory concentrations (22). To minimize the impact of substrate inhibition on KM determination, only the observed reaction rates from 0-20 µM PABP1(437-488) with CARM1 were fit to the Michaelis-Menten equation. There was no statistical difference between either the reported KM 49 or kcat values for CARM1 with PABP1(437-488) (Table 2.1) (p = 0.9540, 0.5629, respectively). The kcat values obtained by FBAPS are within the range of values determined by several groups (47). A recent study analyzed CARM1 methylation of PABP1 peptide and a KM of 12 µM was reported, which is an order of magnitude larger than what is reported here (1.56 µM) (226). It is essential to note that the PABP1 substrate analyzed in this study is larger than the peptide used by Zhang and coworkers, which attributes to the difference between the measured KM values (in addition to different techniques being employed). The lack of difference between FBAPS and the P81 filter binding assay demonstrates the reliability of the assay. Figure 2.3. Michaelis-Menten enzyme kinetics using the FBAPS assay. Comparison of product curves of PRMT1 methylation of GST-GAR (A), and CARM1 methylation of GST-PABP1(437-488) (B) determined through traditional filter binding assay (black circles) or FBAPS (red squares). Error bars represent standard deviation (single experiment performed with three replicates). 50 Table 2.1. Tabulated KM and kcat values comparing FBAPS against filter binding assay. Data from Figure 2.3 were used for calculations. Filter binding assay Enzyme/Substrate KM (µM) kcat (x 10-4 s-1) PRMT1/GAR 1.19 ± 0.2 36.8 ± 1.2 CARM1/PABP1(437-488) 1.54 ± 0.1 6.86 ± 0.2 FBAPS Enzyme/Substrate KM (µM) kcat (x 10-4 s-1) PRMT1/GAR 0.83 ± 0.3 34.6 ± 0.6 CARM1/PABP1(437-488) 1.56 ± 0.6 6.13 ± 1.5 2.3.3 Enzyme inhibition measured by the FBAPS assay We further analyzed the IC50 and KI values of well-characterized inhibitors against PRMT1 and CARM1 by the FBAPS assay (Figure 2.4) to see if they compare against those reported in literature. The KI for SAH against PRMT1 was reported as 6.5 ± 1.6 µM by Thompson and coworkers (227) versus our reported KI of 7.97 ± 2.0 µM (Table 2.2). We see no statistically significant difference (p = 0.2791) between these two values. A similar difference was observed for the KI for SAH against CARM1 where Copeland and coworkers reported 0.16 ± 0.03 µM (99) versus our value of 0.227 ± 0.083 µM (Table 2.2). There is no statistical difference between these values (p = 0.2124). The KI values reported for MS023 against PRMT1 and CARM1 by Eram and coworkers (131) were respectively 11 ± 3 nM and 23 ± 3 nM as compared to our values of 15.4 ± 6.2 nM and 367 ± 98 nM (Table 2.2). The KI values for Diamine 12 and EPZ020411 against PRMT1 and CARM1, respectively, were not reported, so only the apparent IC50 values were compared (132, 133). The IC50 value reported for Diamine 12 (against CARM1) by Therrein and coworkers was 0.2 µM (133) versus our value of 1.97 ± 0.9 µM (Table 2.2). The IC50 value reported for EPZ020411 (against PRMT1) by Mitchell and coworkers was 0.119 µM (132) compared to our value of 0.237 ± 0.02 µM (Table 2.2). 51 The KI values for SAH were the most similar in these comparisons even though they were evaluated using two different techniques (SPA and FBAPS). This similarity can be accredited to the fact that SAH is competing with SAM in both assays (22). When comparing the other values, it is essential to note that the compounds (EPZ020411, Diamine 12, and MS023) all been demonstrated to occupy the peptide binding groove and may display differences in substrate competition depending on the substrate used (131â133). It is for this reason that statistical comparisons (unpaired t-tests) cannot be performed for these compounds. Differences in concentrations of cofactor, substrate, and enzyme, as well as different assays and conditions also contribute to disparate values obtained. Although we see slight differences between our obtained results and those recorded from other assay methods, we see that the FBAPS assay yields comparable results to more expensive techniques. Figure 2.4. Dose-dependent inhibition curves using the FBAPS assay. (A) Chemical structures of inhibitors screened. (B) Inhibition of PRMT1 with EPZ020411 (grey), MS023 (black), and SAH (red), and (C) inhibition of CARM1 with Diamine 12 (blue), MS023 (black), and SAH (red) through the FBAPS assay (the results of three experiment performed on separate days with three replicates each). Error bars represent standard deviation. 52 Table 2.2. Tabulated IC50 and estimated KI values. Data from Figure 2.4 was used for calculations (n = 9). Correlation coefficients (R2) corresponding to nonlinear regressions fitted to equation (2) are indicated. Enzyme/Compound IC50 (µM) Estimated KI (µM) R2 PRMT1/EPZ020411 3.46 ± 0.2 0.237 ± 0.02 0.954 PRMT1/MS023 0.225 ± 0.09 0.0154 ± 0.01 0.968 PRMT1/SAH 140 ± 35 7.97 ± 2.0 0.924 CARM1/Diamine 12 1.97 ± 0.9 0.265 ± 0.06 0.931 CARM1/MS023 2.72 ± 1.4 0.367 ± 0.1 0.942 CARM1/SAH 18.8 ± 6.9 0.227 ± 0.1 0.878 2.3.4 Concluding remarks Hevel and coworkers reported the amenability of using a dot blot with a P81 filter binding assay to increase the throughput of the methylation assay (214), but later reported an incompatibility with protein as opposed to peptide substrates (213). While pre-wetting is an essential step for blotting techniques (228), we found that pre-wetting the P81 membrane prior to applying the reaction was detrimental to the assay and resulted in most of the protein passing through the P81 filter paper onto the Whatman 3MM paper below or into waste. In addition to sample loss, pre-wetting also decreases the resolution of the FBAPS assay as it causes diffusion of proteins and 14C-SAM throughout the membrane, resulting in dot smearing and high background. When PRMT-catalyzed reactions are applied to dry P81 filter paper at low volume (4 µL) under vacuum, the proteins are captured on the filter paper where each spot is distinguishable using Coomassie stain with very little to no methylated protein passing through the filter paper (Figure A6). A unique feature of FBAPS compared to P81 filter binding and scintillation counting is that data can be re-imaged on a storage phosphor screen. Small FBAPS blots can be stored indefinitely with no foreseeable degradation of signal given the 5700-year half-life of 14C (229), whereas storage of scintillation vials for repeated quantification requires a larger footprint and systematic organization of 53 samples. Another advantage of phosphor screening over scintillation counting is the combination of sensitivity (0.9 DPM per mm2 per h) and the dynamic range of 105.4 offered by the phosphor screening function of the Sapphire Biomolecular Imager (230), allowing for quantification of methyltransferase enzymes with weak activity. The limitations with FBAPS are largely represented by limitations with traditional filter binding. The biggest limitation with P81 filter binding paper is the protein binding capacity. We evaluated the maximum protein binding limit by mixing reactions (containing a known amount of methylated GST-GAR) with increasing amounts of non-methylated GST-GAR and spotting onto P81 filter paper (Figure A7). We found that a maximum of 25 µg total protein can be spotted on each dot before detectable loss in storage phosphor signal. Part of this limitation is due to the ionic binding capacity of the P81 filter paper, 18 µeq/cm2 (micro-equivalents of ions able to bind per centimeter squared), that is set by the manufacturer, Thermo Scientific. Another limitation with this technique is pipetting small volumes that can lead to errors in reproducibility. Small volumes were required to improve resolution between spots. Errors associated with reproducibility were overcome by having one experimenter perform all experiments in quadruplicate using the same pipettes. When determining kinetic parameters, it is essential to determine if the substrate to be analyzed will bind the P81 filter paper based on its physicochemical properties. A proteinâs pI and steric size both play a role when in comes to binding filter paper, as it is the number of accessible charged groups that dictate binding (221). Additionally, extreme caution must be taken when analyzing complex mixtures like lysates since other charged proteins will compete for binding the filter paper. It is possible to analyze lysates or greater quantities of protein by using filter paper with a stronger cation exchanger, like LSA-50, which is modified with sulfate groups and has an ionic capacity of 50 µeq/cm2 (VWR). This technique may also be modified to analyze negatively charged substrates by swapping the P81 filter paper for DE81 (Cytiva), which is modified with diethylaminoethyl functional groups. One limitation with this technique is that major vendors like VWR have discontinued the sales of P81 filter paper. There are vendors who prepare phospho-cellulose filter paper that are similar to P81 54 filter paper. St. Vincentâs Institute of Medical Research (https://www.svi.edu.au/resources/phosphocellulose_paper/) sells their custom phospho-cellulose paper that is like P81 but has slight differences in binding capacity. There is also LSA-50 (described above), that is sold by VWR (https://ie.vwr.com/store/product/23804948/ion-exchange-papers) as well as the anion exchange resin, LSB-50, for researchers looking to evaluate the binding of negatively charged analytes. Given the modifications listed, it is possible to adapt this assay to any type of transferase enzyme should the physicochemical properties of the products/substrates allow for their separation by functionalized cellulose filter paper (221). Using P81 filter paper, the FBAPS assay is an adaptation of already existing methods that eliminates the use of scintillation fluid, cuts costs, and increases PRMT assay throughput while yielding comparable results to more expensive technologies. 2.3.5 PRMT2 and filter binding assays The FBAPS assay can measure low levels of radiation and in theory can be used to study enzymes that have lower levels of methyltransferase activity, like PRMT2 (35). Using the same purification protocol described for PRMT1 and CARM1, we purified recombinant human PRMT2 over-expressed in E. coli by Ni-NTA and SEC chromatography to yield >95% pure enzyme (Figure A8). We analyzed the linear range for PRMT2 methylation of RSF1 by filter binding assay (Figure 2.5A) (141). The β-emission resulting from PRMT2 methylation of RSF1 after 10 h was the same magnitude as that obtained for the negative control of RSF1 with radiolabelled SAM. The signal from the negative control from RSF1 and 14C-SAM after 10 h was significantly higher than the signal obtained from the negative controls measured for PRMT1/GST-GAR or CARM1/PABP1 reactions (after 1 or 2 h incubation, respectively). The high background signal from negative controls can likely attributed to the hydrolysis products of SAM, that may be binding to the P81 filter paper or reacting in an uncatalyzed manner with the nucleophilic residues on RSF1 (Figure 2.5B). The hydrolysis of SAM increases with prolonged exposures to elevated temperatures and alkaline conditions, both required for PRMT2 methylation reactions (182). As a result of the overwhelming signal from the no-enzyme control (attributed to 14C-55 SAM degradation products), filter binding assays cannot be used to study PRMT2 or methyltransferase enzymes that require prolonged incubations with 14C-SAM (beyond 2 h) unless the user can distinguish the methylated species of interest (213). Alternative assays are required to study PRMT2 kinetics that can isolate methylated products from SAM degradation products (35). Figure 2.5. Measuring the linear range for PRMT2 methylation of RSF1 by filter binding assay. (A) PRMT2 (1 µM) was incubated with RSF1 (15 µM) and 14C-SAM (50 µM) at 37 ºC from 0-10 h and the reactions were spotted on P81 filter paper and the DPMs were measured by scintillation counting. The dotted line represents the DPMs obtained from the incubation of radiolabelled SAM and RSF1 for 10 h at 37 ºC. Error bars represent standard deviation (single experiment performed with two replicates). (B) Degradation products of [14C-methyl]-S-adenosyl-L-methionine generated by nucleophilic attack from basic groups (e.g., HEPES or amino acids like cysteine) (black arrow), intramolecular carboxylate attack on the γ-carbon (red arrow), primary site of alkaline hydrolysis (blue arrow). The SAM lysis products from the described mechanisms have been colored coordinated and the isotopically labelled carbon indicated. 56 Chapter 3. Development of PRMT2 binding methods1 1 Figure 3.2 presented in this section, and all related results and discussion have been published. Rowley, M. J., Prout-Holm, R. A., Liu, R. W., Hendrickson-Rebizant, T., Ige, O. O., Lakowski, T. M., Frankel, A. (2023) Protein arginine N-methyltransferase 2 (PRMT2) plays a noncatalytic role in the histone methylation activity of PRMT1. Journal of Biological Chemistry 299, 105360. I expressed and purified all recombinant proteins used in this study and performed all activity assays and DSF experiments presented in this chapter. I wrote the entirety of this chapter with editing from Dr. Adam Frankel. 3.1 Introduction The formation of methylarginines is catalyzed by PRMTs (8). PRMTs bind to the methyl cofactor, SAM, and facilitate an SN2 nucleophilic reaction between the electron-rich nitrogen atoms in the guanidino group of arginine residues and the electrophilic methyl sulphonium group to yield methylarginine and SAH (15, 17). While the majority of PRMT enzymes have robust enzymatic activity and can be studied by measuring the accumulation of methylated arginine (using radiolabeled methyl groups or mass spectrometry assays (35, 216, 217, 219)) or SAH (using coupled assays (214, 215, 231)), some PRMTs have poor catalytic activity that cannot be measured using these techniques. PRMT2 is characterized by lackluster in vitro kinetic activity, thus requiring several-hour incubations to yield detectable methylation signal (35). Over long incubation periods, SAM degrades in vitro yielding lower substrate concentrations and an accumulation of SAM degradation products, like SAH, that may inhibit PRMT activity (182, 215). These limitations make it difficult to characterize PRMT2 kinetics in vitro without sensitive techniques that can measure this low activity (35). It has been proposed that PRMT2 exhibits poor kinetic activity since it lacks interaction partners that may be present within cells, or that the substrate against which PRMT2 has optimal activity has yet to be identified (35, 156). PRMT2 has been implicated in several cellular pathways, as a coactivator of transcription, through its purported cellular methylation activity, or via protein-protein interactions (90, 174, 176, 181, 187, 200). Techniques used to interrogate PRMT2 interactions in cellular environments have included, affinity 57 pulldowns, co-IP, yeast-two-hybrid screening, and transactivation reporter assays (90, 165, 174, 176, 187). These cellular-based techniques measure PRMT2 interactions in the presence of other cellular proteins that may be facilitating the reported interactions through complexation with PRMT2 (154, 155, 176, 181, 187). As a result of these drawbacks, it has been challenging to investigate direct PRMT2 interactions with substrates, small molecule inhibitors, or other proteins. There is a demand for the development of non-kinetic in vitro methods that can be used to validate and measure PRMT2 interactions. Given PRMT2 has some capacity for generating methylarginine, the enzyme must bind to SAM and the methyl-accepting substrates (35, 141). When a protein binds to a ligand, it forms a protein-ligand complex stabilized by non-covalent bonds in addition to liberating ordered solvent molecules, making the complex an enthalpically- and/or entropically-favorable interaction (232). These interactions may be accompanied by structural rearrangements of the tertiary structure of a protein, resulting in increased protein thermal stability (233, 234). Changes in protein thermal stability can be measured in vitro using thermal shift assays, such as differential scanning fluorimetry (DSF). In 2017, Cavarelli and coworkers used DSF to demonstrate that both SAH and Cp1 (a synthetic adenosine scaffold connected to a guanidine) could bind to and improve the thermal stability of murine PRMT2 (141). While murine and human PRMT2 are orthologs, there are distinct amino acid differences between these enzymes that may contribute to differences in how the enzymes respond to ligand binding (Figure A9). No study has been performed characterizing the thermal stability of human PRMT2. In this study, we seek to characterize the thermal stability of human PRMT2 by DSF. Using DSF, we evaluated the effects of stabilizing buffer additives on PRMT2 thermal melt to characterize how the protein responds to a range of conditions. We identified conditions that improved PRMT2 thermal stability and performed PRMT2 methylation assays in the presence of these additives to evaluate whether the improved stability led to concomitant improvements in catalytic activity. We additionally examined buffering conditions that have previously been identified to inhibit PRMT2 activity to determine what effects these conditions have on PRMT2 thermal stability. Finally, we examined how five potential 58 ligands interact with human PRMT2 via DSF. Taken together, these data demonstrate it is possible to use DSF to optimize PRMT2 stability and to measure ligand binding to human PRMT2. 3.2 Methods 3.2.1 Reagents SAM was purchased from MilliporeSigma (A7007) and dissolved in 0.5 mM HCl (to prevent hydrolysis) to make a 2.0 mM stock. Sinefungin was purchased from STEMCELL technologies (73182). D-(+)-Trehalose dihydrate was purchased from MilliporeSigma (T9531) and dissolved in H2O to make a 1.82-M stock. L-arginine hydrochloride was purchased from MilliporeSigma (181003) and dissolved in methylation buffer to make a 0.8-M stock. 5000x SYPROâ¢ Orange Protein Gel Stain (in DMSO) was purchased from ThermoFisher (S-6650). Information related to all other reagents has been described in the previous chapter. 3.2.2 Protein expression and purification PRMT2 and GST-GAR expression and purification was described in section 2.2. RSF1 in pET15b was a gift from Vincent Cura. RSF1 was transformed, expressed, and purified following the same protocol as described for PRMT2, with the exception that only a Ni-NTA column was required for purification with no additional SEC column. RSF1 was concentrated in 20 mm Tris/HCl pH 8.0, 100 mm NaCl, and 0.5 mm DTT. 3.2.3 Differential scanning fluorimetry Reactions were prepared in methylation buffer (50 mM HEPES, pH 8.0, 10 mM NaCl, 1.0 mM DTT) with 7.5x SYPROâ¢ Orange to a final volume of 20 µL in a MicroAmp Fast Optical 96-well reaction plate (Applied Biosystems 4346907). Unless otherwise noted, 10 µM PRMT2 was used and enzyme containing reactions were performed in triplicate, while reactions without enzyme were performed in duplicate. To determine the concentration-dependent thermal stability of PRMT2, three PRMT2 concentrations (5, 10, and 15 µM) were evaluated. These concentration-dependent experiments were performed in tandem with glycerol controls where 5 µM PRMT2 was incubated with an equivalent 59 amount of glycerol present with higher concentrations of PRMT2. To determine the effect of osmolytes on PRMT2 thermal stability, reactions were prepared with arginine, glucose, or trehalose concentrations ranging from 0 to 400 mM. To determine the effect of pH on PRMT2 thermal stability, reactions were buffered at pH values ranging from 2 to 8 (citric acid for pH 2.0, acetate buffer for pH 3.0-6.0, and 50 mM HEPES-KOH for pH 6.5-8.0). The effect of PRMT inhibitors on PRMT2 thermal stability was evaluated by incubating 500 µM of either SAM, SAH, Sinefungin, Diamine 12, or 10 µM MS023 (+/- 500 µM SAM). Prepared reaction plates were sealed with an optical adhesive cover (4360954) and incubated in the dark for 30 min at room temperature. Sealed plates were briefly centrifuged at 500 x g using an MPS 100 Microplate Spinner (Labnet). Plates were heated from 25 to 80 ºC at a rate of 0.5 ºC/min and analyzed using the StepOnePlus Real-Time PCR Systems (Applied Biosystems). Samples were excited at 490 nm and emission was measured at 580 nm for 60-s intervals. Data were plotted and analyzed in GraphPad Prism 10.03. To evaluate PRMT melting stability, first-derivative analysis was performed with second order smoothing for data presentation with GraphPad Prism 10.03. The melting temperature, Tm - the steepest point in the fluorescent transition, was derived from generating the second derivative of the raw fluorescence data (no smoothing) and calculating the x-intercept value (using Microsoft Excel 2023) that corresponds to the local maximum on the first-derivative curve. Statistical significance was measured using an ordinary one-way ANOVA test with Bonferroniâs multiple comparison correction applied. A statistical level of significance cut off <0.05 was used. 3.2.4 Gel radiography PRMT2 (1 µM) was incubated with 0.45 mg/mL GST-GAR and 50 or 100 µM 14C-SAM in methylation buffer for 16 h at 37 ºC. The reactions were quenched by addition of SDS loading dye (100 mM Tris-Cl pH 6.8, 0.6% SDS, 10% glycerol, 1.5% BME, 0.02% bromophenol blue) and boiled at 95 ºC for 15 min. Proteins were then separated on Tris/SDS 10% discontinuous PAGE gels. Proteins were stained (in 0.025% blue silver colloidal Coomassie G250), de-stained (10% methanol, 10% glacial acetic 60 acid), and dried down before exposure to a phosphor screen (2 weeks). Phosphor screens were analyzed by phosphor imaging using a Sapphire Biomolecular Imager. 3.3 Results 3.3.1 PRMT2 thermal stability characterization We chose to analyze PRMT2 interactions using DSF, a method that quantifies protein-ligand binding events using a hydrophobic reporter dye, SYPROTM Orange (Figure 3.1 and Tables A1 and A2) (24, 235, 236). In our experiments, we incubated PRMT2 with SYPROTM Orange and measured the dyeâs fluorescent output as a function of temperature (Figure 3.1A). Proteins unfold from thermal stress and reveal hydrophobic patches to which SYPROTM Orange binds, resulting in augmented fluorescence; the melting temperature (Tm) is the steepest point during the unfolding transition, and it is more clearly viewed in a first-derivative plot of a melt curve (24, 235, 236). PRMT2 produced a biphasic melting curve with lower- and higher-temperature melting transitions (Figure 3.1B), Tm1 = 46.4 ± 0.2 ºC and Tm2 = 49.8 ± 0.2 ºC, suggesting two discrete populations that could be constituted by melts of impurities, independently folded domains, or oligomeric populations with distinct thermal stabilities. The observation that PRMT2 exhibited a significant concentration-dependent positive change in Tm values (5 µM PRMT2 Tm1 = 46.4 ± 0.2 ºC and Tm2 = 49.8 ± 0.2 ºC and 15 µM PRMT2 Tm1 =54.5 ± 0.2 ºC and Tm2 = 58.4 ± 0.1 ºC; p-value = 1.01 x 10-10 for ÎTm1 and 1.68 x 10-11 for ÎTm2) points towards accompanying changes to the oligomeric state (Figure 3.1C). When a low concentration of PRMT2 was exposed to increasing amounts of glycerol, some thermal stabilization was observed (5 µM PRMT2 Tm2 =49.8 ± 0.2 ºC and 5 µM PRMT2 + 3% glycerol Tm2 = 54.1 ± 0.3 ºC; ÎTm2 = +4.28 ºC, p-value = 1.45 x 10-5) where Tm2 became the major peak (Figure 3.1D). This glycerol control demonstrates that the glycerol in the PRMT2 storage buffer contributes some thermal stabilization of PRMT2, but there is a greater thermal stabilization when the concentration of PRMT2 enzyme is increased (5 µM PRMT2 and 15 µM PRMT2, ÎTm1 = +8.0, p-value = 1.01 x 10-10, ÎTm2 = +8.5 ºC, p-value = 1.68 x 10-11). The improved PRMT2 stabilization at higher enzyme 61 concentrations can be attributed to both glycerol in the PRMT2 storage buffer and PRMT2 oligomerization. In attempt to measure PRMT2 methylation activity against GST-GAR, we incubated purified his-tagged human PRMT2 with GST-GAR and 14C-SAM for 16 h at 37 ºC and subsequently separated the reactions by SDS-PAGE. The SAM degradation products are not measured when performing gel radiography as they diffuse out of the gel during electrophoresis or during staining/de-staining, allowing the user to specifically monitor for methylated proteins. After exposing the gels for two weeks to phosphor screens (twice the exposure length as previously recorded by Lakowski or Cavarelli (35, 141)), there was no detectable methylation signal above background (Figure A10). We attempted methylation assays of PRMT2 against RSF1, but also failed to generate detectable signal. These data suggest that the human his-tagged PRMT2 used in this study has methylation activity lower than GST-PRMT2 or murine PRMT2. 62 Figure 3.1. PRMT2 thermal stability measured by differential scanning fluorimetry (DSF). Thermal melt of 10 µM PRMT2 in DSF showing (A) averaged raw fluorescence and (B) first derivative data. Evaluation of PRMT2 thermal stability at (C) different protein concentrations, and (D) in presence of increasing concentrations of glycerol at 5 µM PRMT2 (single experiment done in triplicate wells). Raw fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figure A11 and Tables A1âA2. 3.3.2 PRMT2 thermal stabilization by excipients Given the observation that increasing amounts of glycerol thermally stabilized PRMT2 (Figure 3.1D), we sought to explore if other polyols could elicit a greater thermal stabilization upon increased PRMT2 concentration. Polyols are known to influence protein thermal stability through enhanced colloidal stability and have been shown to improve enzyme activity (237â240). We tested the effects of two other polyols, glucose and trehalose, on PRMT2 thermal stability. Both glucose and trehalose resulted in a dose-dependent positive thermal shift of PRMT2, while increasing the height of the first melt transition, Tm1, and decreasing the height of the second melt transition, Tm2 (Figure 3.2A and 3.2B). Glucose and trehalose both shifted the distribution of PRMT2 melting transitions, whereas glycerol 63 caused a condensation of the two melting transitions (Figure 3.2C). At the highest polyol concentration measured (410 mM glycerol, 400 mM glucose, or 400 mM trehalose), glycerol and glucose both imparted higher thermal shifts of PRMT2 (PRMT2 Tm1 = 46.9 ± 0.3 ºC, PRMT2 + 400 mM glucose Tm1 = 51.2 ± 0.7 ºC, ÎTm1 = +4.3 ºC, p-value = 5.96 x 10-9) than trehalose (PRMT2 + 400 mM trehalose Tm1 = 50.0 ± 0.1 ºC, ÎTm1 = +3.8 ºC, p-value = 1.68 x 10-9) (Tables A2, A4, and A6). When we included the polyols in PRMT2 methylation reactions against GST-GAR, we failed to observe any detectable methylation signal via gel radiography and phosphor imaging. Polyols are amongst a group of molecules referred to as osmolytes that can stabilize protein colloidal stability through rearrangement of water molecules around a proteins surface (239, 241, 242). Another osmolyte thought to promote protein colloidal stability is arginine (243â245). Unlike the polyols tested, arginine is zwitterionic and bears a net positive charge at pH 8.0. For our assays, L-arginine HCl was dissolved in water and brought to pH 8.0 (L-arginine HCl dissolved in water is acidic and the low pH solution could mask the arginine effect on PRMT2 thermal stability). We evaluated the effects of adding 0 â 400 mM arginine to the thermal stability of PRMT2, based on literature suggesting that this concentration range is required to observe the osmolyte effect (242). Arginine had no significant effects on PRMT2 thermal stability until concentrations of 100 mM or greater were added (Figure 3.2C). At 100 mM, arginine caused a condensation of the biphasic properties of the PRMT2 melt transition and a significant negative thermal shift compared to PRMT2 Tm2 (PRMT2 Tm2 = 50.8 ± 0.2 ºC, PRMT2 + 100 mM arginine Tm = 48.7 ± 0.1 ºC, ÎTm = -1.4, p-value = 0.016). As arginine concentrations exceeded 100 mM, a dose-dependent decrease in PRMT2 thermal stability was observed until the shape of the melting curves completely changes and we only observe decreased fluorescence with increased temperature. There are reports in literature of arginine acting as a chaotropic agent at high concentrations due to its guanidino group (243). It is highly likely that arginine could be denaturing PRMT2 at concentrations beyond 100 mM, but we did not investigate this possibility. Given the thermal destabilization observed and postulated denaturating effects of arginine, this additive should be avoided in buffers used for PRMT2 as it would be expected to impair PRMT2 interactions (246). 64 Figure 3.2. PRMT2 thermal stability as a function of osmolytes. Influence of increasing concentrations of (A) glucose, (B) trehalose, or (C) arginine on PRMT2 thermal stability (single experiment done in triplicate wells). Raw fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figure A11 and Tables A3âA9. When the Frankel lab first characterized PRMT2 activity in vitro, a series of gel radiography experiments were performed using GST-PRMT2 to methylate GST-GAR while varying salt and pH conditions to identify conditions that afforded highest methylation activity (35). The highest PRMT2 methylation activity was reported with 10 mM NaCl and pH 8.0. It was observed that increasing sodium chloride beyond 10 mM (at pH 8.0) caused a marginal reduction in PRMT2 methylation activity, whereas decreasing the pH of the buffer (with 10 mM NaCl) dramatically decreased the activity of PRMT2. To better characterize PRMT2 biochemistry and the relationship between enzyme activity and thermal stability, we sought to investigate how decreasing pH impacts thermal stability. We measured PRMT2 thermal melt by DSF across decreasing pH conditions (from 8 to 2) (Figure 3.3). To specifically monitor 65 the influence of pH on overall PRMT2 stability, and not oligomeric distribution, we screened these conditions at 10 µM PRMT2 where a single melting population exists. PRMT2 experienced increasingly positive thermal shifts as the pH was decreased from 8 to 5, reaching a Tm of 63.2 ± 0.1 ºC at pH 5, compared to a Tm of 53.2 ± 0.3 ºC at pH 8 (ÎTm = +10.0 ºC, p-value <1.0 x 10-15) (Figure 3.3A and Table A11). When the pH is decreased beyond 5, increasingly negative thermal shifts are observed for PRMT2 (compared to pH 5) until the melting transitions are barely discernable (Figure 3.3B). Extreme pH values are known to denature proteins, which could explain the altered melting curves observed when PRMT2 is exposed to pH of 2, but we did not use any alternative methods to determine whether the protein was aggregated (247). Figure 3.3. PRMT2 thermal stability as a function of pH. Evaluation of PRMT2 thermal stability at (A) neutral to mildly acidic pH (5-8), (B) mildly to highly acidic pH (2-5), and (C) DMSO (single experiment done in triplicate wells). Raw fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figure A12 and Tables A10 and 11. 66 3.3.3 PRMT2 response to small molecules Our PRMT2 binding efforts were focused on well characterized small molecules known to bind either the SAM- or peptide-binding site of PRMTs (131, 133, 141). First, we evaluated how saturating amounts of SAH and SAM (500 µM), the natural ligands of PRMTs, influence PRMT2 thermal stability (Figure 3.4A-B). SAH did not elicit a significant thermal shift of PRMT2 (Tm PRMT2 = 52.8 ± 0.2 ºC, Tm PRMT2 + 500 µM SAH = 53.1 ± 0.1 ºC, ÎTm = +0.4 ºC, p-value = 0.091), while SAM resulted in a significant positive thermal shift (Tm PRMT2 = 53.4 ± 0.1 ºC, Tm PRMT2 + 500 µM SAM = 54.5 ± 0.2 ºC, ÎTm = +1.1 ºC, p-value = 1.4 x 10-5) (Figure 3.4A-B, Tables A12-15). We additionally evaluated sinefungin, a pan-methyltransferase inhibitor, and found that it too evoked significant positive PRMT2 thermal shift (Tm PRMT2 = 53.4 ± 0.1 ºC, Tm PRMT2 + 500 µM sinefungin = 54.6 ± 0.1 ºC, ÎTm = +1.2 ºC, p-value = 8.2 x 10-6) (Figure 3.4C and Table A14). Sinefungin is a chemical analog of SAM where a protonated primary amine extending from the 5â ribose ring (Figure 3.4C) mimics the methyl sulphonium structure in SAM (Figure 3.4C), potentially explaining why these molecules stabilize PRMT2 in a similar manner not seen by SAH. It is plausible that SAM and sinefungin could have a higher affinity binding interaction with PRMT2 than SAH. The CARM1-specific small molecule inhibitor, Diamine 12, failed to influence PRMT2 thermal stability (Tm PRMT2 = 53.4 ± 0.1 ºC, Tm PRMT2 + 500 µM Diamine 12 = 53.7 ± 0.1 ºC, ÎTm = +0.2 ºC, p-value = 0.13) (Figure 3.4D and Table A14). We additionally tested MS023 (10 µM) in the presence or absence of saturating SAH against PRMT2 and noted a small but statistically significant stabilization (within a single experiment) when both MS023 and SAH were present (Tm PRMT2 = 52.8 ± 0.2 ºC, Tm PRMT2 + 500 µM SAM + 10 µM MS023 = 53.3 ± 0.2 ºC, ÎTm = +0.5 ºC, p-value = 0.078) (Figure 3.4E and Table A15). Previous studies have shown that MS023 binds strongest to PRMTs when bound with SAH (24, 131), hence why we measured both conditions. When we incubated the 0.5% DMSO control (for MS023) with PRMT2, we observed a similar small stabilization that is not statistically different from PRMT2 incubated with MS023 and SAH (Table A15), indicating that thermal stabilization in this case is driven by DMSO rather than ligand-induced stabilization (Tm PRMT2 + 0.5% DMSO = 53.3 ± 0.1 ºC, Tm PRMT2 + 500 µM SAM + 10 µM MS023 = 53.3 ± 0.2 ºC, 67 ÎTm = -0.01 ºC, p-value = >0.99). Taken together, these results suggest that only SAM and sinefungin bind PRMT2 in a thermally stabilizing manner at the concentrations measured, and we failed to demonstrate PRMT2 thermal stabilization by SAH, MS023, and Diamine 12. Figure 3.4. PRMT2 stabilization by PRMT small molecule probes. Evaluation of PRMT2 thermal stability in presence of (A) SAH, (B) SAM, (C) Sinefungin, (D) Diamine 12, and (E) MS023 (+/- SAH) all at 500 µM (single experiment done in triplicate wells). Thermal melt of PRMT2 in presence of small molecule inhibitors shown in black, and in the absence of small molecules is shown in cyan. Chemical structures of the small molecules shown to the right of the first-derivative plot. Raw fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figure A13 and Tables A12âA15. 3.4 Discussion 68 3.4.1 PRMT2 thermal melt and activity PRMT2 produced a biphasic melt curve by DSF. Proteins displaying multiphasic melting properties in DSF may indicate the presence of impurities, independently-folded domains, or oligomeric populations with distinct thermal stabilities (24, 248, 249). In this study, we found that human PRMT2 produced a largely biphasic melt depending on protein concentrations and additives. Since the more thermally stable population (Tm2) became dominant in the presence of increased protein concentration, we postulated it was due to concentration-dependent PRMT2 homo-oligomerization, common among PRMTs (23). The PRMT2 thermal stabilization observed with glycerol can be attributed to the osmotropic effect of glycerol on colloidal protein stability (237). The utility of DSF is limited in that it does not allow for any assessment of the oligomerization state of each melting population. It is important to note that DSF is entirely dependent on protein dye interactions and the results obtained by DSF should be corroborated with an additional assay where the readout is dye independent. The lack of reported methylation activity for hexa-histidine-tagged human PRMT2 was unexpected since similar conditions were used that yielded PRMT2 activity as a GST-tag or hexa-histidine-tagged murine PRMT2 (35, 141). The difference in reported activity between murine and human PRMT2 could potentially be a result of the difference in sequence between the two orthologs, namely in the N-termini of each protein (Figure A9). Côté and coworkers identified several PRMT1 splice variants with variable truncations about the N-terminus that had varying degrees of activity (250). No such study has been conducted on the N-terminus of PRMT2 but based on the sequence similarity between PRMT1 and PRMT2, the N-terminus of PRMT2 may also play a vital role in catalytic activity. It is plausible that the GST-tag could have had an impact on PRMT2 activity, hence why there was activity observed for GST-PRMT2 during the 2009 study and not in the subsequent study (35). Upon comparing the purity of PRMT2 used in this study against the GST-PRMT2 used in the 2009 study, there were several contaminants present in the GST-PRMT2 that might have contributed to the improved PRMT2 activity (35). 3.4.2 Exploring PRMT2 thermal stability with solution additives 69 In attempt to better characterize the biphasic distribution of PRMT2, we investigated how osmolytes influence PRMT2 thermal stability. Osmolytes are small organic molecules that promote protein thermal stability by lowering the free energy of folded proteins through a combination of entropic and enthalpic contributions, depending on the osmolyte (241). Polyols are highly hydrophilic molecules that are thought to exert their protein stabilization effects through ordering water molecules in a solution, reducing the entropic penalty for protein folding, and binding polar side chains (237, 239, 244). Glycerol is a polyol present in PRMT2 storage buffer as a cryoprotectant, that also appears to protect PRMT2 from thermal stress. The thermal protection afforded by glycerol towards PRMT2 was also observed by the larger polyols, glucose and trehalose, similar to that seen in previous studies investigating the thermally protective properties of glycans (237, 239). Interestingly, glycerol caused a condensation of the biphasic melting properties of PRMT2, while glucose and trehalose caused a change in the distribution of the two melting transitions of PRMT2. We postulate that longer polyols could be stabilizing the lower order PRMT2 oligomeric state, composing the first melting transition, while glycerol could be stabilizing the higher order PRMT2 oligomeric states. Once methods are established to characterize the distribution of PRMT2 oligomeric states in solution, it will be pertinent to evaluate how the polyols impact the distribution of PRMT2 oligomeric states. Since the polyols failed to improve PRMT2 catalytic activity while eliciting a thermally stabilizing response, it is unlikely that lack of PRMT2 stability can be attributed as the reason for PRMT2 catalytic inactivity. Amino acids are another group of osmolytes capable of improving protein colloidal stability (241). Arginine is unique amongst the osmolytes since its effect on protein stability is specific to each protein, as opposed to the global stabilization observed by polyols (241, 242, 245). PRMT2 was thermally destabilized and possibly denatured in presence of increasing concentration of arginine. Based on literature, we predict that at low arginine concentrations, the presence of an extra cationic charge (from the guanidino group of arginine) caused enhanced electrostatic repulsion between PRMT2 monomers, explaining the negative thermal shift seen at low arginine concentrations. At higher arginine concentrations, we predict the guanidine on arginine displaces water to bind the amide backbones on 70 PRMT2, leading to unfolding of the enzyme (like urea-based denaturation) (243, 244). Arginine could be used to study PRMT2 unfolding or as a substitute for urea in denaturing PRMT2 purifications, but not to promote aggregation. To explore conditions that inhibit PRMT2 activity, we examined the relationship between decreasing solution pH and PRMT2 thermal stability. In the absence of excipients and synthetic compounds, we observed the greatest PRMT2 thermal stability when the pH was equal to the isoelectric point of PRMT2 (pI = 5.03). At the isoelectric point, PRMT2 net charge is zero and there will be the least electrostatic repulsions between protein monomers (237, 251). While highly thermodynamically favorable, it has been previously demonstrated that when the pH of solutions buffering PRMT2 methylation reactions is decreased below 8, PRMT2 catalytic activity also decreases (35). Net positive charge on PRMT2 appears to be important for enzymatic activity, potentially providing some insights into substrate binding or catalytic activity. Perhaps the basic patches and overall positive charge on PRMT2 could provide a docking platform for negatively charged substrates, like nucleosomes. This presents the first instance of using DSF to evaluate the effects of excipients on the thermal stability of a PRMT enzyme. While the excipients did not improve PRMT2 responsiveness to ligands, they did improve overall thermal stability and yield monophasic melt transitions. A study conducted in 2016 demonstrated how DSF can be used to identify buffer conditions that promote protein folding (246). The so-called DSF-guided refolding, or DGR, identified thermally stabilizing buffering conditions that facilitated chemical refolding of several enzymes. Our findings indicate that osmolytes are an important class of molecules that improve protein thermal stability that should be included in DGR experiments. This method could be an asset to research laboratories attempting to refold proteins, improve the thermal melt of a protein for ligand binding, or to identify optimal activity conditions for human his-tagged PRMT2. 3.4.3 PRMT2 interactions with PRMT inhibitors 71 Several groups have demonstrated that PRMT2 can bind SAM either using binding assays, or indirectly through methylation assays (35, 90, 174, 194). Here we show that both SAM and sinefungin result in positive thermal shifts for PRMT2, indicating stabilizing interactions. PRMT1 also experienced a positive thermal shift in presence of both of these ligands (24). Upon closer comparison, it can be noted that the magnitude of thermal stabilization experienced by PRMT2 in response to either SAM or sinefungin is 5-fold lower than the stabilizations observed when PRMT1 was exposed to either ligand (24). PRMT1 and PRMT2 share 31.52% sequence identity, while adopting a similar structural fold. It is not surprising given the differences between the two proteins that there are stark differences in how they respond to SAM and sinefungin. Unlike PRMT1 and murine PRMT2 (141), human PRMT2 did not display thermal stabilization in the presence of SAH. A close comparison of human and murine PRMT2 sequences (Figure A9) reveal differences between residues in the cofactor binding domain, the SH3 domain, and the dimerization arm that may account for any thermal stability differences in response to SAH. Another possibility may be attributed to differences in the DSF experiment itself (e.g., different buffers and SAH concentrations used). Of particular concern is the high concentration of SAH (1 mM) used by Cavarelli and coworkers against PRMT2. Such high concentrations of SAH are not biologically relevant and could result in non-specific interactions, potentially as an osmolyte. Unfortunately, the authors investigating murine PRMT2 did not present the complete thermal shift data and therefore a direct comparison cannot be made between murine and human PRMT2 melts. The difference in stabilization of PRMT2 by SAM and SAH could be indicative of a structural feature present in the human PRMT2 SAM binding site that is sensitive to the chemistry about the sulfur atom of SAM/SAH. Validation of this hypothesis would require the structure of human PRMT2 to be solved in complex with SAM or sinefungin. Based on the thermal stabilization of PRMT2 through binding SAM or sinefungin, future studies focused on the development of small molecule inhibitors against PRMT2 should include chemical features that mimic SAM or sinefungin. While there has been direct evidence in literature demonstrating PRMT2 can bind SAM, the same cannot be said for MS023. Several groups have claimed that MS023 can inhibit PRMT2 function in a 72 cellular context, but failed to demonstrate direct evidence of an interaction between MS023 and PRMT2 (185, 186). In this work, we failed to demonstrate an interaction between PRMT2 and MS023 via DSF. Previous studies have demonstrated that SAH-binding significantly strengthens the interaction between PRMTs and MS023, but this was not the case for PRMT2 (24, 131). It is essential that an orthogonal assay be performed to validate whether PRMT2 can interact with MS023, such as isothermal titration calorimetry (ITC) or surface plasmon resonance (SPR). Unlike PRMT2, MS023 binding and/or inhibition to PRMT1, CARM1, PRMT6, and PRMT8 has been characterized by biophysical experiments (131). The authors who employed MS023 to inhibit arginine methylation by PRMT2 (in GBM and HCC cells lines) failed to account for the inhibition of all other Type I PRMTs and did not demonstrate a PRMT2 interaction with MS023 (185, 186). To demonstrate MS023 binding to PRMT2 in cells, the cellular thermal shift assay (CETSA) could be evaluated. CETSA evaluates how ligand binding protects protein from thermal stress in cells (252, 253). CETSA could be used to validate a PRMT2 interaction with MS023 in cells, should PRMT2 aggregate in cells when exposed to thermal stress (252). 73 Chapter 4. Protein arginine N-methyltransferase 2 (PRMT2) plays a noncatalytic role in the histone methylation activity of PRMT11 1 A version of chapter 4 has been accepted for publication with the Journal of Biological Chemistry. Rowley, M. J., Prout-Holm, R. A., Liu, R. W., Hendrickson-Rebizant, T., Ige, O. O., Lakowski, T. M., Frankel, A. (2023) Protein arginine N-methyltransferase 2 (PRMT2) plays a noncatalytic role in the histone methylation activity of PRMT1. Journal of Biological Chemistry 299, 105360 I expressed and purified all recombinant proteins used in this study, performed all in vitro methylation assays and was responsible for the enzyme kinetics studies. Riley Austin Prout-Holm performed cloning on the PRMT2 SH3 domain into pET28a(+). Rui Wen Liu performed mutagenesis on PRMT2 to generate the PRMT2H112Q mutant. Thordur Hendrickson-Rebizant and Olufola O. Ige expressed and purified recombinant Xenopus laevis histone proteins used in preliminary experiments of this study. Dr. Ted M. Lakowski generated the GST-PRMT2ÎSH3, His6CARM1, His6-PRMT6 clones used in this study and contributed to editing the manuscript and experimental design. Dr. Adam Frankel and I designed experiments, analyzed data, and wrote and edited the manuscripts. 4.1 Introduction The histone code is comprised of a multitude of post-translational modifications (PTMs) on histone proteins that control access to the genetic code and determine cell fate (96). Epigenetic marks constituting the histone code are deposited by a plethora of protein-modifying enzymes, often referred to as the writers of the epigenetic code (254, 255). These marks alter the biochemical properties of histone proteins and require specific reader proteins to allow transcriptional machinery to access the underlying genes (96). Aberrant expression of epigenetic writers can lead to increased expression of oncogenic genes and repression of tumor suppressor genes that can contribute to the onset of cancer (75, 255). Understanding the writers of the epigenetic code is of the utmost importance for developing targeted cancer therapeutics to prevent uncontrollable cell growth (75). Amongst the epigenetic writers, within the protein arginine N-methyltransferase (PRMT) family of enzymes (8, 10, 17), PRMT2 remains somewhat enigmatic. We and others have shown that it is able to 74 catalyze the formation of monomethylarginine (MMA) and asymmetric dimethylarginine (aDMA) on histone H4, as well as on glycine- and arginine-rich, and serine- and arginine-rich substrates using S-adenosyl-L-methionine (SAM) as the methyl donor that gets converted into S-adenosyl-L-homocysteine (SAH) after methyl transfer (35, 141). However, PRMT2 is distinct from enzymes that form either the same products (PRMT1, PRMT3, CARM1, PRMT6, and PRMT8), MMA only (PRMT7), or MMA and symmetric dimethylarginine (sDMA) (PRMT5 and PRMT9), in that it exhibits the lowest catalytic efficiency for methyl transfer (47). It is conceivable that we have yet to identify substrates upon which human PRMT2 exhibits high activity. Despite such low in vitro activity, paradoxically PRMT2 was shown to facilitate aDMA formation on histone H3 at arginine 8 (H3R8me2a) at promoter sites for transcriptional activation during Xenopus development (181). It was later demonstrated that PRMT2 overexpression and the H3R8me2a mark were linked to oncogenic transcriptional programming in glioblastoma multiforme (GBM), hepatocellular carcinoma (HCC), and renal cell carcinoma (RCC), in each case strongly correlating with disease severity (184â186). In this regard, PRMT2 joins the PRMT pantheon of other well-known writers with oncogenic capacity (58, 124). These other writers include transcriptional activators PRMT1 responsible for H4R3me2a, and CARM1 responsible for H3R17me2a, H3R26me2a, and H3R42me2a (108, 109, 112, 113), and transcriptional repressors PRMT5 responsible for H2AR3me2s, H3R8me2s, and H4R3me2s, and PRMT6 responsible for H3R2me2a (116, 117, 137, 212, 256). Like other PRMTs, the PRMT2 monomer possesses a SAM-binding Rossmann fold, a peptide binding groove, and dimerization arm, and it forms a toroidal shape as a homodimer comprised of catalytic cores (PDB ID: 5FUL) (141, 194). Unlike other PRMTs, PRMT2 contains an N-terminal SH3 attached to the rest of PRMT2 through a highly flexible linker. This auxiliary SH3 domain has been shown to bind proline-rich motifs on proteins involved in splicing and cell scaffolding (30, 187, 200). The importance of the SH3 domain for PRMT2 function was initially highlighted in a study showing that its removal resulted in an interaction loss between the methylation substrate hnRNP E1B-AP5 and its 75 ostensible methyltransferase PRMT2 (HRMT1L1) in cultured H1299 cells (176). In activity assays, removal of the SH3 domain was shown to further reduce the already low catalytic activity of PRMT2 (35, 141). While not linked to enzymatic activity per se, full-length PRMT2 has been shown to function as a transcriptional coactivator of several nuclear receptors (174), as well as influence functions of Iï«B-a (165), and the retinoblastoma gene product (167). It is unclear with our present understanding of PRMT2 if it functions in cells as an active methyltransferase, as a structural subunit to facilitate methylation by another more active subunit as we have observed with PRMT1 (19, 35), or a combination of both. In this study, we sought to better understand how PRMT2 interacts with histones and other PRMT enzymes to determine whether any alternative mechanisms of PRMT2-mediated histone methylation exist. We used thermal shift assays to evaluate whether PRMT2 can bind to histone peptides and to assess heteromeric PRMT2 protein-protein interactions. These thermodynamic findings led us to identify in vitro conditions where PRMT1/2 interact (19, 257), which we confirmed by using fluorescently-tagged PRMT1/2 enzymes and native PAGE analysis. We complemented these techniques by demonstrating PRMT2âs influence on the catalytic activity of PRMT1 towards calf thymus histones, independent histones, histone octamers, and mononucleosomes. Our findings demonstrate that PRMT2 enhances PRMT1 catalytic activity towards histone substrates. 4.2 Methods 4.2.1 Materials SAM, SAH, and lyophilized calf thymus histones were purchased from Millipore Sigma (A7007, A9383, and H9250, respectively) and dissolved in 0.5 mM HCl to make 2 mM stock (SAM, SAH) or water to make 0.6 mg/mL stocks (histones). Radiolabeled cofactor S-adenosyl-L-[methyl-14C]-methionine (14C-SAM; 58 mCi mmol-1) was purchased from PerkinElmer (NEC363050UC). The H4 (Ac-SGRGKGGKGLGKGGAKR) peptide was purchased from Canada Peptide and was synthesized with N-terminal acetylation. The H3 (Ac-ARTKQTARKSTGGKAPRKQLA) and H2A (Ac-SGRGKQGGKARAKAKTRSSR) peptides were purchased from Biomatik and synthesized with N-76 terminal acetylation. Peptides were dissolved in water to prepare 2 mM stocks. Recombinant human H3.3 mononucleosomes and H3.3 octamers were purchased from EpiCypher (16-0012 and 16-8012, respectively). SYPROâ¢ Orange Protein Gel Stain was purchased from ThermoFisher (S-6650) as a 5000x concentrate in dimethylsulfoxide (DMSO) and was diluted to a 50x stock in water immediately before use. NativeMarkTM unstained protein standards were purchased from Invitrogen (LC0725). 4.2.2 DNA constructs Cloning of PRMT1 (UnitProt ID: Q99873-3), PRMT1E153Q, PRMT2 (UnitProt ID: P55345-1), PRMT2E220Q, mCer-PRMT1, and mCit-PRMT2 in pET28a(+) (Novagene) have been previously described (19, 163). Cloning of PRMT2âSH3 (aa 83-433) in pGEX-2T (Novagene) has been previously described (35). PRMT2H112Q and the PRMT2 N-terminus (aa. 1-103) were generated by mutagenesis (Quick Change Mutagenesis Kit, Agilent) on the PRMT2-pET28a(+) template using either: 5â- CGT GGC AGG ATG AAG AGT ACT TCT AAG GCA GCT ATG GAA CTC TG-3â and 5â- CAG AGT TCC ATA GCT GCC TTA GAA GTA CTC TTC ATC CTG CCA CG-3â primers for PRMT2 SH3 that inserted a stop codon after the SH3 domain (aa 1-103), or 5â- AGC TAT GGA ACT CTG AAA CTC CAG TTG GAG ATG TTG GCA GAC CAG-3â and 5â- CTG GTC TGC CAA CAT CTC CAA CTG GAG TTT CAG AGT TCC ATA GCT-3â for PRMT2H112Q. Pet28a_Synthetic_Human_H2A.1 (UniProt ID: P0C0S8) and Pet28a_Human_H3.3 (UniProt ID: P84243) were gifts from Joe Landry (Addgene plasmids # 42634 and # 42632, respectively). pMMTH+H1.4 (UniProt ID: A3R0T8) plasmid was a gift from Michael Rossen (Addgene plasmid # 157796). cePRMT5 in pET21a(+), and PRMT3, PRMT7, and PRMT8 in pGEX-2T were generous gifts from Rui-Ming Xu (Chinese Academy of Sciences) and Steven Clarke (UCLA), respectively. Human CARM1 in pET28a(+) has been described elsewhere (223). pET3+H3 was a gift from Dr. Karolin Luger (Colorado State University). Histone H2A mutants were generated by mutagenesis (Quick Change Mutagenesis Kit, Agilent) on the Pet28a_Synthetic_Human_H2A.1 (+) template using: 5â â GAG ATA TAC CAT GTC TGG TAA GGG TAA ACA AGG TGG TAA AGC C â 3â and 5â â GGC TTT ACC ACC TTG TTT ACC CTT ACC AGA CAT GGT ATA TCT C â 3â primers for H2AR3K; 5â â GTG GTA AAC AAG GTG GTA AAG 77 CCA AGG CCA AAG CCA AAA CCC GCT â 3â and 5â â AGC GGG TTT TGG CTT TGG CCT TGG CTT TAC CAC CTT GTT TAC CAC â 3â primers for H2AR11K; 5â â GGT CTG CAA TTT CCG GTC GGT AAC GTT CAT CGT CTG CTG CGC AAA â 3â and 5â â TTT GCG CAG CAG ACG ATG AAC GTT ACC GAC CGG AAA TTG CAG ACC â 3â primers for H2AR29N. Histone H3 mutants were generated by mutagenesis (Quick Change Mutagenesis Kit, Agilent) on the Pet28a_Human_H3.3 template using: 5â - ACA AAG CAG ACT GCC GCC AAA TCG ACC GGT GGT â 3â, and 5â - ACC ACC GGT CGA TTT GGC GGC AGT CTG CTT TGT â 3â primers for H3R8A. 4.2.3 Protein expression and purification All PRMT2 constructs were transformed and expressed in Escherichia coli Arctic Express (DE3) (Stratagene), except for GST-PRMT2âSH3, which was expressed in E. coli BL21 DE3 pLysS. PRMT8 was transformed in E. coli BL21(DE3) pLysS (Stratagene). Histone H2A was transformed into E. coli Arctic Express DE3 (Stratagene), whereas histones H3 and H4 were transformed in E. coli Rosetta DE3 (Stratagene). In brief, cells were grown in LB media to an OD600 of 0.6 and protein expression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) at 16 ºC for 16 h. Cells were harvested via centrifugation (10000 x g, 4 ºC, 15 min) and cell pellets were frozen at -80ºC. Purity of enzymes used in this chapter was assessed by SDS-PAGE analysis (Figure A14). 4.2.4 His-tagged PRMT purification Cell pellets containing hexa-histidine tagged proteins were thawed on ice in a lysis buffer [2 mL/g wet weight of cells; 50 mM HEPES-KOH pH 7.4, 1 M NH4Cl, 10 mM MgCl2, 10 mM imidazole, 0.1% Triton X-100, 0.1% lysozyme, 25 U mL-1 DNAse I, 1.0 mM phenylmethanesulphonyl fluoride (PMSF), 7 mM β-mercaptoethanol (BME), and 1.0 mM EDTA-free protease inhibitor cocktail (PIC)] and incubated for 1 h at 25 ºC before further lysis via freeze-thaw. Homogenized cell lysates were immersed in liquid nitrogen until frozen, followed by immersion in a 25 ºC water bath until fully thawed and repeated for a total of three cycles. Soluble proteins were separated by centrifugation (35000 x g, 4 ºC, 1 h) and filtered through a 0.22 µM low protein binding PVDF membrane (Millex). Clarified lysates containing hexa-histidine tags were applied to pre-equilibrated Ni-NTA resin or HisTrap FF column (GE 78 healthcare) in wash buffer [50 mM HEPES-KOH pH 7.6, 1 M NH4Cl, 10 mM MgCl2, 10 mM BME, 10 mM imidazole, 1 mM PMSF] and eluted using stepwise imidazole elution buffers or over a gradient of 50 â 400 mM imidazole when performing column elution (elution buffer had same composition as wash buffer except for 100-400 mM imidazole). Column purification performed using AKTA FPLC basic and protein elution monitored by A280 UV-Vis trace. Fractions containing the protein (confirmed by 10% SDS-PAGE) were applied to a pre-equilibrated HiLoad26/600 Superdex 200 pg size-exclusion column (SEC) in wash buffer [50 mM Tris-HCl, 100 mM NaCl, 1 mM DTT, pH 8.0]. Eluted fractions containing the desired protein (evaluated by 10% SDS-PAGE analysis) were pooled and concentrated in a storage buffer [hexa-histidine tagged proteins: 100 mM HEPES-KOH pH 8.0, 200 mM NaCl, 1 mM DTT, 10% glycerol, 2 mM EDTA; GST-tagged proteins: 50 mM Tris-HCl pH 7.5; 500 mM NaCl; 1 mM DTT, 10% glycerol (v/v)] using Amicon Ultra 15 mL filters [30-kDa molecular weight cut-off (Millipore)]. The fluorescently tagged proteins were quantified spectrophotometrically according to the extinction coefficient of the fluorophore: mCit (ε516 nm = 77000 M-1 cm-1) and mCer (ε434 nm = 43000 M-1 cm-1). All other protein concentrations were quantified by the Bradford assay (Quick Startâ¢ Bradford Protein Assay; BioRad). 4.2.5 GST-tagged PRMT purification Cell pellets containing GST-tagged proteins (PRMT3, PRMT7, PRMT8, and PRMT2ÎSH3) were lysed using the same protocol as listed in the main text except using a PBS lysis buffer [2 mL/g pellet; 1x PBS pH 7.3, 5% glycerol; 1 mM EDTA; 1 mM EGTA, 0.1% Triton X-100, 0.1% lysozyme, 25 U mL-1 DNAse I, 1.0 mM phenylmethanesulphonyl fluoride (PMSF), 7 mM β-mercaptoethanol (BME), and 1.0 mM EDTA-free protease inhibitor cocktail (PIC)]. Following separation of soluble proteins, GST-tagged proteins were injected to a pre-equilibrated 5 mL GST-Trap FF column (GE healthcare) (pre-equilibrated in PBS pH 7.3 and 1 mM DTT) and eluted with reduced glutathione (50 mM Tris-HCl, 10 mM reduced glutathione, 1 mM DTT, pH 8). Protein concentration and quantification was performed using same protocols described previously. 79 The GST-tag was cleaved from PRMT2ÎSH3 by incubating the protein with thrombin in PBS pH 7.3 for 16 h. The uncleaved GST-tagged protein, free GST-tag, and thrombin was captured on the GST-Trap FF column and 1 mL Benzamidine FF column (GE healthcare) (pre-equilibrated in PBS pH 7.3, 1 mM DTT) and the cleaved protein was collected in the flow-through. 4.2.6 Histone purification X. laevis histone H4 (same amino acid sequence as human histone H4) was purified according to previously published methods (19), whereas human histones H2A and H3 were purified according to the rapid histone purification method (258). Cell pellets containing native histones (H2A and H3 constructs) were thawed on ice in a lysis buffer [5 mL/g pellet; 6 M Urea, 40 mM sodium acetate pH 5.2, 200 mM NaCl, 10 mM lysine, 0.1% Triton X-100, 1.0 mM PMSF, 5 mM BME, and 1.0 mM EDTA-free protease inhibitor cocktail (PIC)] and incubated for 1 h at 25 ºC before further lysis via freeze-thaw. Homogenized cell lysates were immersed in liquid nitrogen until frozen, followed by immersion in a 25 ºC water bath until fully thawed and repeated for a total of three cycles. Soluble proteins were separated by centrifugation (35000 x g, 4 ºC, 1 h) and filtered through a 0.22-µM low protein binding PVDF membrane (Millex). Clarified lysates were applied to pre-equilibrated QHP column coupled to an SP-HP column (GE healthcare) in wash buffer [6 M Urea, 40 mM sodium acetate pH 5.2, 10 mM lysine, 5 mM BME, 1 mM PMSF]. After the lysate was applied and 1 column volume (CV) was passed, the QHP containing the DNA was removed and the column was washed with another 4 CV of wash buffer. The bound histones were eluted over a NaCl gradient (same composition as wash buffer except for 1 M NaCl). Protein fractions were separated on a 16.5% Tricine-SDS-PAGE to confirm which fraction contained the protein. Eluted proteins were placed in 5000-Da MWCO dialysis tubing and dialyzed against refolding buffer [100 mM HEPES-KOH pH 8.0, 2 M NaCl, 1 mM BME, 1 mM EDTA] over 12 h at 4 ºC. The dialysate was replaced twice to ensure removal of all salts. Histones were quantified by spectrophotometry according to the extinction coefficient, H2A and H3.3 ε280 = 4470 M-1cm-1. 80 4.2.7 Differential scanning fluorimetry Reactions were prepared in methylation buffer (50 mM HEPES, pH 8.0, 10 mM NaCl, 1.0 mM DTT) with 7.5x SYPROâ¢ Orange to a final volume of 20 µL in a MicroAmp Fast Optical 96-well reaction plate (Applied Biosystems 4346907). Unless otherwise noted, 5 µM PRMT2, PRMT2âSH3, or PRMT2H112Q was used, and all reactions were performed in triplicate. To determine effect of peptide and cofactor on PRMT2 stability, reactions were prepared with SAH, H2A, H3, or H4 peptide from 0-500 µM in triplicate. To determine heteromeric protein stabilization effects of PRMT1 towards PRMT2, PRMT2âSH3, or PRMT2H112Q, either 1, 5, or 10 µM PRMT1 were incubated with 1, 5, or 10 µM PRMT2 (except for PRMT2H112Q and PRMT2âSH3 where only 5 µM were tested). All running parameters and analysis methods have been described in aim 4. All running parameters and analysis methods have been described in Chapter 3. To evaluate PRMT2âSH3 Tm, the raw data were normalized as previously described (24). The normalized data were fit to a Boltzmann Sigmoidal curve using equation (1) from which the Tm could be generated. (1) ð¦ = ðµðð¡ð¡ðð + (ðððâðµðð¡ð¡ðð)1+ð(ððâð¥ððððð) KD values for specific ligands were generated by plotting Tm values against the concentration of ligand as a dose-response curve. The curve was fit to a single site ligand binding model using equation (2) where x is the ligand concentration, y is the Tm, and P is the protein concentration. (2) ð¦ = ðµðð¡ð¡ðð + (ððð â ðµðð¡ð¡ðð)(1 âðâð¾ð·âð¥+â(ð+ð¥+ð¾ð·)2â4ðð¥2ð) 4.2.8 Native PAGE PRMT enzymes (1 µM each) were mixed with methylation buffer and incubated at 37 ºC for 1 h before being mixed with native PAGE electrophoresis loading buffer (100 mM Tris-Cl pH 6.8, 10% glycerol, 1.5% BME, 0.02% bromophenol blue) and applied to a 4-20% gradient PAGE gel. Gels were run with 5 µL NativeMarkTM unstained protein standards for molecular weight (MW) estimation. The unstained gels were analyzed for absorbance at 365 nm using a BioRad Gel Doc XR imager using trans-81 UV setting with 0.1-s exposure (to detect mCerulean). Fluorescence of mCitrine-tagged proteins was detected by exciting the unstained gels at 488 nm and measuring emission at 520 nm using a Sapphire Biomolecular Imager. Proteins were stained and de-stained as described previously. 4.2.9 Histone methylation kinetics PRMT1 (100 nM) was pre-incubated with PRMT2 (0, 100, 1000, or 10000 nM) at 37 ËC for 1 h before addition of 0-25 µM histone H2A or H3 and 10 µM 14C-SAM for an additional incubation at 37 ËC for 2 h. The reactions were spotted (2 x 2 µL) in triplicate on P81 filter paper via dot-blot apparatus under vacuum. The filter paper was washed, stained, and radioactivity quantified by FBAPS (259). The relative phosphor signals were plotted against H2A, H3, or H4 protein concentrations, and the data were fit to either a simple linear regression model or the Michaelis-Menten equation (3) using Graph Pad Prism 9.5.0 to derive apparent values KM (App) and kcat (App). (3) ð£ =ðððð¡ (ð´ðð)[ð¸]ð[ð]ð¾ð (ð´ðð)+[ð] 4.2.10 Gel radiography Unless indicated otherwise, PRMT2 (or PRMT2 variant) was titrated from 10-10,000 nM against 2 µM of either PRMT1, 3, 4, 5 (C. elegans PRMT5, incubations at 25 ºC), 6, 7, or 8 in methylation buffer and incubated for 1 h at 37 ºC to allow for complexation. The methylation reactions were initiated by addition of 0.6 mg/mL calf thymus histones and 10 µM 14C-SAM, and further incubated an additional 2 h. The reactions were quenched by addition of tricine-SDS loading dye (50 mM Tris-Cl pH 6.8, 0.32% SDS, 13% glycerol, 0.4% BME, 0.02% bromophenol blue) and boiled at 95 ºC for 15 min. Proteins were then separated on Tris/Tricine/SDS 16.5% discontinuous PAGE gels. Proteins were stained, de-stained (described above), and dried down before exposure to a phosphor screen (exposure times are indicated in figure legends). Phosphor screens were analyzed by phosphor imaging using a Sapphire Biomolecular Imager. Densitometry analysis was performed by Image Studio Lite Ver 5.2, as described previously (259) to generate relative changes in histone methylation. 82 4.3 Results 4.3.1 PRMT2 ligand binding characterization Our binding efforts were focused on ligands described in previous studies indicating they may bind directly to PRMT2: SAH (35), histone H2A peptide (260), histone H3 peptide (181), and H4 peptide (35). We monitored both PRMT2 melting populations (i.e., Tm1 and Tm2) over a broad range of ligand concentrations (Figure 4.1, Figure A15, and Tables A16 and A17). We noted some batch-to-batch variability in PRMT2 response due to different glycerol percentages per stock protein concentration, but this issue did not appear to affect relative thermal shifts. SAH failed to elicit thermally stabilizing responses towards either of the PRMT2 Tm values (Figure 4.1A). All histone peptides caused a dose-dependent condensation of the two melting populations into a single Tm with concomitant thermal stabilization (Figure 4.1B-1D). The histone peptides exhibited melting characteristics with SYPRO OrangeTM independent of PRMT2 (all peptides yielded 10-fold lower fluorescence and indistinct Tm values without PRMT2). Hence, we conclude thermal stabilization observed was due to the interaction between PRMT2 and histone peptide (Figure A15). Compared to the apo state, H2A, H3, and H4 peptides at 500 µM each elicited ÎTm of +2.0 ºC, +7.8 ºC, and +5.9 ºC for ligand bound PRMT2, respectively. By analyzing Tm1 of PRMT2 in the presence of histone H2A and H3 peptides, DSF-derived KD values of 145 and 35 µM were determined (Figure 4.1B and 1C). A KD for histone H4 peptide binding to PRMT2 could not be generated due to poor model fitting (Figure 4.1D). We additionally tested whether PRMT2 could bind both SAH and H3 peptide together by DSF (Figure A16 and Tables A18 and 19). Compared to PRMT2 with H3 peptide, no change in Tm was seen, but the first-derivative peak height increased, indicating a steeper melt transition for the complex. 83 84 Figure 4.1. Effects of ligand binding on PRMT2 thermal stability. PRMT2 was incubated with increasing amounts of (A) SAH, (B) H2A peptide, (C) H3 peptide, or (D) H4 peptide for DSF measurements. First-derivative melting curves (in relative florescence units per second (RFU/s)) are shown on the left while notable Tm trends for each of the ligands with PRMT2 are shown on the right. DSF-derived KD was determined for H3 peptide binding to PRMT2. Data presented are averages of three replicates conducted during a single experiment; error bars show standard deviations. Raw fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figure A15 and Tables A16 and A17. Given the lack of responsive from PRMT2 to SAH via DSF, we sought alternative approaches to evaluate whether PRMT2âs interaction with SAH is important for the histone peptide thermal stabilization. We made a PRMT2 SAM-binding mutation, H112Q, that has previously been shown to inhibit PRMT2 catalytic activity in cells (186). Like native PRMT2, PRMT2H112Q also displayed clearly biphasic melting properties, but the second melting transition was too broad to accurately define a melting temperature (Figure A17A). The PRMT2H112Q Tm1 was 2.4 ºC lower than PRMT2 Tm1 and the melt was overall much lower in raw fluorescence with gradual changes in fluorescence compared to the steeper transitions observed for wildtype PRMT2 (Figure A17B). When PRMT2H112Q was incubated with either SAH and/or H3 peptide (saturating amounts), there was no statistically significant change in melting transition observed compared to the apo state (Tables A20 and A21). No KD measurement was performed for PRMT2H112Q with H3 peptide due to the lack of responsiveness at saturating H3 peptide. 4.3.2 SH3 domain effects on PRMT2 thermal stabilization To determine whether the PRMT2 SH3 domain played a role in binding to the histone H3 peptide, we performed DSF on the PRMT2 construct cloned without the N-terminus containing the SH3 domain (PRMT2ÎSH3). Unlike the full-length construct, PRMT2ÎSH3 produced a monophasic melt curve with very gradual changes in fluorescence (Figure A18A and Tables A22-24). This broad fluorescence change made differentiation of data difficult to interpret; therefore, data were normalized and fit to a Boltzmann sigmoidal nonlinear regression equation in which the Tm is the temperature value at 50% of maximum fluorescence (Figure A18B and A18C). The Tm obtained for PRMT2ÎSH3 was 39.5 ºC, 7.0 ºC lower than full-length PRMT2 Tm1 at 46.5 ºC (Figure A17D and Tables A22-24). When incubated with histone H3 peptide, PRMT2ÎSH3 was stabilized by 2.6 ºC with 500 µM H3 peptide (no 85 significant shift below this concentration) compared to 7.8 ºC stabilization for full-length PRMT2 with 500 µM H3 peptide. Like full-length PRMT2, SAH negligibly impacted the melt curve of PRMT2ÎSH3 either in the presence or absence of H3 peptide. A KD greater than 1400 µM was estimated from Tm values for the H3 peptide binding to PRMT2ÎSH3 (Figure A18E and Tables A22-24). To further validate that the PRMT2 SH3 domain can bind to H3 peptide, we attempted DSF on the PRMT2 N-terminus, containing the SH3 domain (without the PRMT scaffold, amino acids 1-103). Unfortunately, there were no fluorescence differences between the SYPRO OrangeTM dye alone and dye with PRMT2 N-terminus (measured at multiple concentrations ranging from 100 nM to 100 µM data not shown), indicating that DSF could not be used to measure peptide binding to the PRMT2 N-terminus. We attempted isothermal titration calorimetry on the PRMT2 N-terminus with both H3 and H4 peptides and observed negligible changes in heat of binding (data not shown). 4.3.3 PRMT1/2 complex response to ligands Our group previously demonstrated that PRMT2 can bind to PRMT1 in vitro using co-IP and in cells using BiFC, as well as increase PRMT1 activity in vitro towards histone H4R3, suggesting that these two enzymes can interact (19). We sought to investigate whether the PRMT1/2 complex can be thermodynamically characterized by DSF. We conducted control experiments on PRMT1 in the absence of PRMT2 to evaluate how it interacts with the H3 peptide with or without SAH. PRMT1 showed a small thermal stabilization in presence of 500 µM H3 peptide and a much greater increase in stabilization when H3 peptide and SAH were present at 500 µM each (Figure A19). We determined a DSF-derived KD for H3 peptide binding to PRMT1 of 105 µM in the presence of 500 µM SAH (Figure A19 and Tables A25-27). We used DSF on PRMT1 with PRMT2 (PRMT1/2), PRMT2ÎSH3 (PRMT1/2Î), and PRMT2H112Q (PRMT1/2H112Q) to probe for conditions that change protein thermal stability indicative of protein-protein interactions (Figure 4.2, Figures A20-22, and Tables A28-33) (261, 262). In the absence of SAM and H3 peptide, the mixed thermal stability of PRMT1/2 and PRMT1 alone showed no differences, and the PRMT1/2Î mixture yielded a biphasic melt with the major peak between the melting 86 temperatures of PRMT1 and PRMT2ÎSH3 (Figure 4.2B). In contrast, the PRMT1/2H112Q mixture yielded a biphasic melt with a small but evident shoulder (Tm1) that was 1.6 ºC lower than PRMT2H112Q and the major peak (Tm2) that was 0.9 ºC higher than PRMT1 Tm1 (Figure 4.2B). In the presence of saturating SAH concentration, we saw no difference between the Tm of the PRMT1/2 population and PRMT1 (Figure 4.2C and Table A29). The thermal melt of the PRMT1/2Î mixed population with SAH had two distinct melting temperatures, where Tm1 was close to PRMT1 with SAH, but Tm2 was greater than the Tm values of each individual PRMT melt with SAH (Figure 4.2C and Table A31). The PRMT1/2H112Q mixture yielded biphasic curves in presence of saturating amounts of either SAH or H3 peptide, neither peak showed significant differences in thermal stability compared to the individual PRMT melts (Figure 4.2C-D and Table A33). The addition of a saturating H3 peptide concentration to PRMT1/2 or PRMT1/2Î yielded two distinct peaks in the first-derivative plot that mimicked peaks seen for each enzyme alone with H3 peptide (Figure 4.2D and Table A30). In Figure 4.2E, both H3 peptide and SAH incubated with PRMT1/2 resulted in a single sharp peak representative of a steep monophasic melt transition that appeared to combine Tm values from individual PRMTs. In the absence of the SH3 domain, however, the peak height of PRMT1/2Î with both ligands was not as sharp and with a lower Tm than what is seen for PRMT1/2, suggesting that the SH3 domain adds to complex stability in response to H3 peptide and SAH (Figure 4.2E and Table A31). The PRMT1/2H112Q mixture in presence of saturating H3 peptide and SAH yet again yielded a biphasic melting transition, but Tm1 was 1.7 ºC more stable than PRMT2H112Q, and Tm2 was 1.5 ºC less stable than PRMT1 (Figure 4.2E and Table A33). We additionally evaluated how varying the concentration of PRMT1 (1-10 µM) while holding PRMT2 constant at 10 µM and vice versa impacts the changes in melting transitions in the different conditions listed above (Figure A23 and Table A34). We found that PRMT1 and PRMT2 influenced each otherâs thermal stabilities primarily in the presence of both SAH and H3 peptide. 87 Figure 4.2. PRMT1/2 response to ligands by DSF. (A) Modular schematics depicting the three PRMT2 variants evaluated in complex with PRMT1. The PRMT2ÎSH3 protein contains a 102-amino acid deletion of the PRMT2 N-terminus. PRMT2H112Q has a single point mutation at position 112 (marked with a purple line). First-derivative melting curves (in relative florescence units per second (RFU/s)) of PRMT1 and PRMT2 (left), PRMT2ÎSH3 (middle), or PRMT2H112Q (right), was evaluated by DSF (B) in the absence of ligands, (C) presence of 500 µM SAH, (D) presence of 500 µM H3 peptide, or (E) presence of both SAH and H3 peptide at 500 µM each (single experiment done in triplicate wells). Raw 88 fluorescence spectra, tabulated melting temperatures, and statistical evaluation available in Figures A20-22 and Tables A30-35. The evidence that PRMT1 and PRMT2 can influence one anotherâs thermal stability in a cofactor- and domain-dependent manner, prompted us to determine whether a PRMT1/2 interaction could be observed with mCerulean (mCer) and mCitrine (mCit) fluorescent fusions of PRMT1 and PRMT2 using native PAGE (Figure 4.3). We used substrate concentrations above KI and KM values for PRMT1 towards SAH and H3 peptide, respectively (47). In the absence of SAH or H3 peptide, both mCer-PRMT1 and mCit-PRMT2 displayed multiple banding patterns consistent with monomeric, dimeric, and higher order oligomers with no overlap between mCer or mCit signals when the two enzymes were mixed. The addition of SAH caused mCer-PRMT1 bands to condense into one single band at ~450 kDa, producing an increase in mCer signal compared to the apo state, whereas no mCit-PRMT2 banding pattern change was apparent upon SAH addition. The H3 peptide consolidated the higher fluorescent bands for mCit-PRMT2 to a uniform species around ~600 kDa while also reducing the signal for the lower molecular weight species regardless of PRMT1 presence. The opposite is seen for mCer-PRMT1, where the H3 peptide caused a reduction in the highest band by Coomassie stain, and an emergence of more signal at ~70 and 150 kDa. When the fluorescent populations were mixed with H3 peptide, the emergence of a single band containing mCer and mCit emission was observed in the overlay (Figure 4.3, lane 13). We saw a more pronounced, consolidated band bearing both mCer-PRMT1 and mCit-PRMT2 in the presence of both SAH and H3 peptide (Figure 4.3, lane 18). These results mimic the same DSF responsiveness to ligands for PRMT1 and PRMT2 individually and in combination (Figure 4.2). 89 Figure 4.3. Native PAGE gels of fluorescent PRMT1/2 heteromeric complexes. PRMTs each at 1 µM were incubated for 1 h at 37 ºC in the presence or absence of SAH (10 µM) and/or histone H3 peptide (200 µM). The proteins were separated by electrophoresis on a 4-20% native PAGE gel and imaged to detect fluorescent proteins at their specific excitation wavelengths. The overlay is a composite of fluorescent images. 90 4.3.4 PRMT2 impact on PRMT1 substrate specificity To assess whether the interaction between PRMT1 and PRMT2 impacts PRMT1 enzymatic activity, we evaluated the methylation activity of PRMT1/2 (at varying molar ratios) towards calf thymus histones using 14C-SAM (Figure 4.4, Figures A24-26, and Table A35). Calf thymus histones were used as an initial screening substrate given its low cost, ease of sample preparation, and presence of multiple arginine-rich proteins. To visualize and distinguish methylation of the individual histones, we separated reactions by gel electrophoresis on 16.5% Tricine-SDS-PAGE gels that were subsequently stained with Coomassie blue to visualize proteins, and the dried gels were exposed to phosphor screens to quantify 14C-methylated substrate. Our first experiment was similar to our previous study, where we incubated 100 nM PRMT1 with increasing amounts of PRMT2 (from 10-10000 nM) and monitored histone H4 methylation, but here we used calf thymus histones (Figure 4.4A). In the absence of PRMT2, PRMT1 methylation activity towards histone H4 was barely detectable, and PRMT2 activity at 10 µM was not detectable. With increasing PRMT2 concentrations in the presence of PRMT1, the H4 methylation signal concomitantly increased up to 5 µM PRMT2; higher PRMT2 concentrations resulted in a decreased H4 methylation (Figure 4.4A, lanes 12 and 13). Detectable levels of histone H2A methylation can also be seen in the phosphor image. We performed a control experiment with a titration of PRMT2 storage buffer and observed decreased H4 methylation at high glycerol concentrations, indicating increased H2A and H4 methylation is PRMT2-specific (Figure A24). We repeated the same experiment with 10-fold more PRMT1 (1 µM), while keeping all other variables constant (Figure 4.4B). At this concentration, PRMT1 alone predominantly methylated histone H4 but also methylated histones H3 and H2A to a lesser extent (Figure 4.4B, lane 3). With increasing PRMT2 against 1 µM PRMT1, histone H4 methylation remained the same while histone H2A methylation dramatically increased and histone H3 methylation decreased (Figure 4.4B, lanes 4-13). These results demonstrate that PRMT2 can influence PRMT1 activity in a concentration-dependent manner. For this reason, we refer to this experiment as the âinfluencer assay.â We performed control 91 reactions with a catalytically inactive PRMT1E153Q and observed no histone methylation, even in presence of PRMT2, confirming that histone methylation is dependent on PRMT1 activity (Figure A25). To further probe the mechanism of how PRMT2 influences PRMT1 activity, we performed the influencer assay on PRMT1 with PRMT2 mutants. The SAM-binding mutant PRMT2H112Q did not show any effect on PRMT1 methylation of calf thymus histones (Figure 4.4C), demonstrating that the ability of PRMT2 to bind SAM is critical for how it influences PRMT1 activity. PRMT2E220Q, previously demonstrated to inhibit PRMT2 methylation activity (19), showed the exact same pattern of enhanced H2A methylation as did wildtype PRMT2 (Figure 4.4D). Thus, PRMT2 catalytic activity is not required for it to influence PRMT1 methylation of histones H4 and H2A. The two truncations of PRMT2 we tested were the PRMT2 N-terminus and PRMT2 without its SH3 domain (PRMT2ÎSH3). When we performed the influencer assay using the PRMT2 N-terminus against PRMT1, we saw a dose-dependent increase in the methylation of both histones H2A and H4 (Figure 4.4E), indicating that the PRMT2 N-terminus can augment PRMT1 activity towards select histone substrates. Without the SH3 domain, PRMT2ÎSH3 failed to improve PRMT1 activity above the activity of PRMT1 alone (Figure 4.4F). We sought to evaluate how different ratios of PRMT1/2 (1:0, 1:1, 1:10, 1:100) are able methylate each of the calf thymus histones as a function of increasing substrate concentrations (Figure 4.4G-4.4I). This was done by performing densitometry on the phosphor images from the gels (Figure A26 and Table A35). At highest PRMT2 concentrations (1 and 10 µM) and lowest calf thymus histone concentrations, PRMT1 methylated PRMT2 to a similar extent as histone H4 from calf thymus histone, but PRMT2 methylation was inhibited at higher histone concentrations (Figure A26). Only after a 10-fold PRMT2 excess of PRMT1 did we see noticeable changes in the histone methylation pattern, specifically a three-fold increase in the methylation signal of histone H2A at 50 µg/mL calf thymus histones (Figure 4.4G). At a 100-fold PRMT2 excess, the increase in histone H2A methylation signal occurred after 100 µg/mL (Figure 4.4G), histone H3 methylation signal decreased over all substrate concentrations compared to other PRMT1/2 ratios (Figure 4.4H), and the highest substrate concentrations were required to yield 92 histone H4 methylation signals achieved with lower PRMT1/2 ratios (Figure 4.4I). Due to the heterogenous nature of calf thymus histones purified from eukaryotic cells and bearing post-translational modifications, we were unable to use this data to derive any quantitative enzymatic properties from these gels. 93 94 Figure 4.4. PRMT1/2 methylation of calf thymus histones. (A) PRMT1 (100 nM) or (B) PRMT1 (1 µM) methylation reactions using 10 µM 14C-SAM and 10 µg calf thymus histones with increasing amounts of PRMT2. Panels C-F are the same conditions as described in panel B with the following PRMT2 variants: (C) PRMT2H112Q, (D) PRMT2E220Q, (E) PRMT2 N-terminus, (F) PRMT2ÎSH3. Reactions were separated on a 16.5% Tricine SDS-PAGE and visualized by Coomassie staining (top), followed by phosphor imaging analysis after a 24-h exposure (bottom). Phosphor signal from PRMT1 (100 nM) methylation towards histone (G) H2A, (H) H3, or (I) H4 as a function of increasing calf thymus histone amount in presence of increasing amounts of PRMT2 (0, 100, 1000, 10000 nM). Raw data for panels G-I are available in Figure A26 and Table A35. Our next set of experiments were designed to evaluate how PRMT2 can influence PRMT1 activity towards purified and unmodified histone proteins in different macromolecular contexts. We expressed and purified histones H2A, H3, and H4 for in vitro PRMT1/2 methylation analysis. In these experiments, we titrated the protein substrate against different molar ratios of PRMT1:PRMT2 (1:0, 1:1, 1:10, 1:100) and measured methyl transfer by the filter binding and phosphor screening (FBAPS) (259) for individual histones (Figures A27-29). Densitometry data for reactions are presented after subtracting phosphor signal from the histone-only control. We observed no detectable signal for reactions with PRMT2 alone (Figure A30). Without PRMT2, PRMT1 methylation activity towards histone H2A was best fit to a simple linear regression, indicating the KM (App) could be beyond the concentrations tested (Figure 4.5A). In the presence of 1:1 and 1:10 PRMT1/2 ratios, H2A methylation data fit to a nonlinear regression model (Michaelis-Menten) where the KM (App) decreased to 128 and 44.7 µM at 1:1 and 1:10 PRMT1/2 ratios, respectively (Table 4.1). The total methylation signal for PRMT1 towards H2A increased by several fold at 1:1 and 1:10 PRMT1/2 ratios compared to PRMT1 alone (Figure 4.5A and Figure A27). Based on the Michaelis-Menten fit, there is a dose-dependent decrease in kcat (App) for the PRMT1 methylation of H2A as the PRMT1/2 ratio increased above 1:1 (Table 4.1). A similar trend in kcat (App) increase was also observed when PRMT2 was added to PRMT1 methylation reactions against histone H3, except for the 1:100 ratio where methylation appeared inhibited by PRMT2 (Figure 4.5B, Table 4.1 and Figure A28). The increase in kcat for PRMT1 methylation of H3 with increasing PRMT2 is also accompanied by a dose-dependent increase in KM for 1:1 and 1:10 ratios (Figure 4.5B, Table 4.1). Accordingly, the apparent specificity constant kcat/KM (App) remained relativity constant for histone H2A with PRMT1/2 at ratios of 1:1 to 1:10 but increased nearly 5-fold at the 1:100 ratio. In contrast, kcat/KM 95 (App) for histone H3 was similar with PRMT1 only and the 1:1 presence of PRMT2 but decreased at least 2.7-fold at a 1:10 PRMT1/2 ratio. PRMT1 methylation of histone H4 in the absence of PRMT2 had the highest kcat (App) of all the substrates tested, as expected based on the gel radiography and previous studies (Figure 4.5C, Table 4.1) (19, 47, 263). As PRMT2 was introduced to the PRMT1 methylation reactions with histone H4, there was an increase in phosphor signal (Figure 4.5C and Figure A29). With increasing PRMT2, there was a trend of decreasing KM (App) values while the kcat (App) remains relatively consistent, but the data fit poorly to the Michaelis-Menten model (Figure 4.5C, Table 4.1). We attempted to measure the activity of PRMT1/2 towards lower histone H4 concentrations to determine whether PRMT2 can decrease the PRMT1 KM (App) for histone H4, but unfortunately PRMT1 methylation of PRMT2 dominated the signal (like that observed in Figure A26). Given the data presented for histone H4, the kcat/KM (App) for histone H4 increased roughly by 2- and 6-fold at 1:1 and 1:10 PRMT1/2 ratios above PRMT1 alone, but a different method is required to evaluate nanomolar histone H4 methylation reactions to obtain accurate fits (Table 4.1). 96 Figure 4.5. PRMT1/2 methylation of histones in different macromolecular contexts. PRMT1 (100 nM) methylation reactions (2 h) with 0, 100, 1000, or 10000 nM PRMT2 towards recombinant histones (A) H2A, (B) H3, and (C) H4 using 10 µM 14C-SAM and signal quantification by FBAPS (single experiment done in triplicate). (D-E) PRMT1 (100 nM) and/or PRMT2 (1 µM) methylation reactions (2 h) towards recombinant histone H2A mutants using 10 µM 14C-SAM. (F) PRMT1 (1 µM) and/or PRMT2 (1 µM) methylation reactions (2 h) towards recombinant histone H3 mutants using 10 µM 14C-SAM. (G) PRMT1 (100 nM) or (H) PRMT1 (1 µM) methylation reactions (16 h) using 10 µM 14C-SAM and 2.5 µM recombinant H3.3 histone octamer with increasing amounts of PRMT2 (12-h exposure for phosphor image). (I) PRMT1 (1 µM) methylation reactions (16 h) using 10 µM 14C-SAM and 1 µM recombinant 97 H3.3 mononucleosomes with increasing amounts of PRMT2 (1-week exposure for phosphor image). Reactions were separated on a 16.5% Tricine SDS-PAGE and visualized by Coomassie staining (top), followed by phosphor imaging analysis (bottom). Raw data for panels A-C and densitometry for panel E available in Tables A36-38 and Table A39, respectively.98 Table 4.1. Tabulated apparent kinetic parameters for PRMT1/2 methylation of histones H2A, H3, and H4. Data from Figure 4.5A-C were used for calculations. The data for the PRMT1 (PRMT1/2 ratio 1:0) methylation of H2A, PRMT1/2 (1:100) methylation of H3, and PRMT1/2 (1:100) methylation of H4 were fit by simple linear regression analysis yielding the following lines of best fit: H2A: y = 0.01461x + 7.733 (R2 = 0.941); H3: y = 96.59x + 4273 (R2 = 0.501); H4 y = 1055x + 32391 (R2 = 0.546), respectively. Adjusted correlation coefficients (R2) corresponding to nonlinear regressions fit to equation (3) for H2A, H3, and H4 data are indicated. PRMT1/2 Ratio PRMT1 PRMT1/2 (1:1) PRMT1/2 (1:10) PRMT1/2 (1:100) PRMT1 PRMT1/2 (1:1) PRMT1/2 (1:10) PRMT1/2 (1:100) PRMT1 PRMT1/2 (1:1) PRMT1/2 (1:10) PRMT1/2 (1:100) Substrate H2A H3 H4 kcat (App) (x 10-6 s-1) n/a 26.6 ± 7.4 10.7 ± 1.7 0.480 ± 0.03 1.02 ± 0.04 5.44 ± 0.4 10.9 ± 0.4 n/a 267 ± 170 79.8 ± 22 76.2 ± 4.0 n/a KM (App) (x 10-6 M) n/a 128 ± 37 44.7 ± 6.5 0.456 ± 0.2 0.992 ± 0.10 4.02 ± 0.4 28.2 ± 12 n/a 38.8 ± 14 6.22 ± 3.4 1.66 ± 01.0 n/a kcat/KM (App) (M-1s-1) n/a 0.208 ± 0.08 0.239 ± 0.05 1.05 ± 0.5 1.03 ± 0.11 1.35 ± 0.2 0.387 ± 0.2 n/a 6.87 ± 5.0 12.8 ± 7.9 33.7 ± 20 n/a R2 n/a 0.962 0.805 0.519 0.802 0.848 0.955 n/a 0.961 0.718 0.380 n/a 99 Our next aim was to determine whether PRMT2 changes the site of methylation for PRMT1 on histones. We previously demonstrated that H4R3 is the primary site of augmented arginine methylation by PRMT1/2 showing that the H4R3K peptide was unable to be methylated (19). Using site directed mutagenesis, we made the following histone N-terminal tail mutations: H3R8A, H2AR3K, H2AR11K, and H2AR29N. These purified proteins were assayed to determine the major methylation site for PRMT1/2. We evaluated the activity of PRMT1, PRMT2, and PRMT1/2 against wildtype and mutant versions of these histones by gel radiography to visualize changes in activity (Figure 4.5D-E). Compared to wildtype histone H2A, H2AR3K resulted in the greatest reduction in activity of the three mutants by either PRMT1 or PRMT1/2 (Figure 4.5E and Table A39). Although H2AR3K showed reduced methylation, some residual signal was detectable, indicating that there could be low-level methylation at other sites. H2AR11K also decreased methylation compared to wildtype H2A, although to a lesser extent than H2AR3K (Figure 4.5E and Table A39). When we screened the histone H3R8A mutation against PRMT1 and PRMT1/2, there was a ten-fold reduction in methylation compared to wildtype histone H3, indicating R8 is the primary arginine methylated by PRMT1 and PRMT1/2 (Figure 4.5F). In order to present histone substrates in a more biologically relevant context, we tested the PRMT2 influencer assay on different histone complexes. We performed exhaustive methylation reactions with 100 nM or 1 µM PRMT1 at different PRMT2 concentrations against recombinant histone H3.3 octamers (Figure 4.5G-H). At both PRMT1 concentrations, PRMT2 appeared to elicit an increase in PRMT1 activity towards histone H4, and to a lesser degree other lower molecular weight contaminants. In contrast, the exhaustive methylation of unmodified recombinant nucleosomes with 1 µM PRMT1 revealed a robust PRMT2-dependent increase in the methylation of histone H2A and H4, as well as methylation of an unknown partial degradation product that migrated just below H2A (Figure 4.5I). At 10 µM PRMT2 with 1 µM PRMT1, an increased methylation of histone H2B was also detected. These results indicate that PRMT2 influences the amount of PRMT1 methylation activity and histone substrate specificity within a recombinant nucleosome substrate context. 100 To determine whether the alteration in substrate specificity induced by PRMT2 towards PRMT1 was a unique phenomenon for this pairing, we repeated the same PRMT2 titration against PRMT3-8 with calf thymus histones as the substrate (Figure 4.6, Table 4.2). Amongst all combinations, high concentrations of PRMT2 inhibited histone H3 and H4 methylation (except for PRMT1). When PRMT2 was titrated against CARM1, we observed modest increases in the methylation of a band at ~32 kDa consistent with the size of histone H1 (Figure 4.6B). Reactions with PRMT6 showed a PRMT2 dose-dependent increase in histone H2A methylation (Figure 4.6D). These results suggest that PRMT2 can influence other PRMTs and cause changes to histone methylation. 101 Figure 4.6. PRMT2 influence on PRMT methylation of histones. (A) Crude PRMT3 (48-h exposure), (B) CARM1 (48-h exposure), (C) C. elegans PRMT5 (cePRMT5) (2-week exposure), (D) PRMT6 (24-h exposure), (E) crude PRMT7 (48-h exposure), and (F) PRMT8 (24-h exposure) (1 µg each) methylation reactions using 10 µM 14C-SAM and 10 µg calf thymus histones with increasing amounts of PRMT2. (E) Reactions were separated on a 16.5% Tricine SDS-PAGE and visualized by Coomassie staining (top), followed by phosphor imaging analysis (bottom). Exposure time for each phosphor image indicated next to corresponding enzyme used. 102 Table 4.2. PRMT methylation activity towards calf thymus histones +/- PRMT2. Recombinant PRMT enzymes were purified to varying extents and pre-complexed with PRMT2 for 1 h at 37 ºC (except cePRMT5, which was incubated at 25 ºC) prior to a 2-h incubation with 10 µg calf thymus histone and 10 µM 14C-SAM at 37 ºC. Proteins were separated on 16.5% tricine-SDS-PAGE gel, visualized by Coomassie stain, and methyl transfer was analyzed by phosphor imaging (exposure times available in Figure 20). The reported methylation patterns are relative to the respective gel/enzyme. Enzyme No PRMT2 PRMT2 (10 µM) H1 H2A/H2B H3 H4 H1 H2A/H2B H3 H4 PRMT1 - + + +++ - +++ - +++ PRMT3 - + - ++ - + - + PRMT4 - + +++ - + - + - cePRMT5 - - - ++ - - - - PRMT6 - + ++ +++ - +++ - ++ PRMT7 - - + ++ - - - ++ PRMT8 - + - +++ - - - + 4.4 Discussion 4.4.1 PRMT heteromeric complexation The formation of PRMT heteromeric complexes is not a novel observation. The first investigations into interactions between PRMT2 and other PRMTs employed transient expression of green fluorescent protein (GFP) fusions of PRMT1-8 in HEK293 cells stably expressing GFP-PRMT2 (39). Immunoprecipitation (IP) of PRMT2 and Western blotting against GFP showed that PRMT2 was able to pull down GFP-PRMT8, which had previously been shown to be mediated between PRMT2âs SH3 domain and two proline-rich stretches in PRMT8âs N-terminus (211), as well as faint bands consistent with GFP-PRMT1 and GFP-PRMT6. Our group identified permissible conditions for formation of a PRMT1/6 complex in vitro using GFP variants (mCerulean and mCitrine), which were used to quantitate PRMT hetero- and homo-oligomerization by FRET analysis (23). A recent study added CARM1 to the list of PRMT enzymes capable of forming heteromeric complexes with other PRMT enzymes (154). Using IP techniques, these authors demonstrated that PRMT2 and CARM1 can pull down one another from MCF7 cells along with the BRD4 protein (154). It was further shown that CARM1 methylation of BRD4 was influenced by PRMT2, as upregulation of PRMT2 increased its methylation, and sgRNA silencing of PRMT2 led to decreased methylation (154). In parallel with the discovery of the PRMT2/4 complex, another group identified PRMT5 and PRMT6 as functionally associating enzymes as 103 oncogenic drivers in colorectal cancer (CRC) (153). This growing body of evidence challenges the view of PRMT enzymes as solo writers of the methylarginine mark and instead recognizes that these enzymes can act in concert to modulate their methylation activities. Our group investigated heteromeric PRMT1/2 interactions both in vitro and in HeLa cells using co-IP, biomolecular fluorescence complementation (BiFC), and enzymatic assays (19). PRMT1/2 co-immunoprecipitated one another in vitro and in HeLa cells. BiFC showed that PRMT1/2 co-localized in HeLa cells in an SH3 domain-dependent fashion (19). Our enzymatic assays demonstrated enhanced PRMT1 activity towards recombinant histone H4 in vitro in a PRMT2 concentration-dependent fashion and a global increase in MMA, aDMA, and sDMA levels on total cellular protein with ectopic expression of PRMT1 and PRMT2 in HeLa cells (19). Our findings led us to propose that PRMT2 can form a heteromeric complex with PRMT1 and possibly other PRMTs. In this study, we used DSF, native gel electrophoresis with fluorescently tagged enzymes, and activity assays to identify conditions where PRMT1/2 form favorable interactions. DSF has been previously used to study protein complex formation by comparing the thermal stability of a complexed state to that of a non-complexed state (261, 262). Our group previously established conditions for measuring cofactor and peptide ligand binding to human PRMT1 in vitro using the DSF assay (24), and Cavarelliâs group performed ligand binding studies on murine PRMT2 via DSF, revealing thermal stabilization of the protein at 1-mM concentrations of SAH, sinefungin, and the bisubstrate analogue Cp1 relative to the apo protein (141). 4.4.2 Human PRMT2 thermal stability characterization PRMT2 melts varied considerably in response to different ligands. A novel finding for this study was that histone peptides representative of the N-terminal tails of histone H2A, H3, and H4, all could elicit a thermally stabilizing response with PRMT2 (Figure 4.1). The peptide binding preference for PRMT2 was H3 > H2A > H4 via DSF. We have previously established an important mechanistic relationship between cofactor binding (SAM or SAH) and dimer formation (23), which did not appear to be the case for PRMT2 here. Due to the lack of responsiveness from PRMT2 to SAH in the presence or 104 absence of histone H3 peptide, we cannot comment on the binding order of PRMT2 to its substrates using DSF. In lieu of a functional assay for measuring PRMT2 interaction with SAH, we evaluated how PRMT2H112Q, the PRMT2 SAM-binding mutation, could interact with ligands (186). The PRMT2H112Q completely inhibited the interaction between PRMT2 and histone peptides, indicating that the ability to bind to cofactor is required for peptide substrate binding. 4.4.3 Non-canonical SH3 domain functions The PRMT2 SH3 domain retains the same vital structural elements of a canonical SH3 domain, yet its amino acid sequence does not align well with sequences of other SH3 domains (192, 193). Several studies have demonstrated that the PRMT2 SH3 domain is capable of binding to canonical polyproline stretches, despite differences in sequence alignment (30, 187). Our DSF and methylation data suggest that the PRMT2 SH3 domain facilitates histone protein binding. Akin to the PRMT2 SH3 domain, the Mona Gads SH3 domain is capable of binding to both proline-rich as well as an R/KXXK motif where X is any amino acid (193, 264). We aligned SH3 domain structures of PRMT2 and Mona Gads complexed to its peptide ligand and speculate that basic residues within the R/KXXR/K motifs found in all four core histones can bind to the acidic regions of the PRMT2 SH3 domain (Figure A31). Given our data for the PRMT2 and PRMT2ÎSH3, it appears that the SH3 domain contributes to the binding of PRMT2 to the histone peptides, but due to a lack of evidence demonstrating direct peptide binding, we cannot draw any conclusions from our results in this study. Alternative methods for measuring SH3 binding to these basic peptides will need to be investigated to identify a binding mechanism. The role of the PRMT2 SH3 domain has remained elusive. Splice variants of PRMT2 bearing the SH3 domain and the cofactor-binding domain without an intact dimerization arm and beta-barrel domain (i.e., C-terminal truncations) have been implicated in breast cancer (160, 161).These variants were shown to be capable of augmenting estrogen receptor alpha-mediated transactivation activity. Yet other studies have shown that the SH3 domain and much of the N-terminal portion of PRMT2 was not required for nuclear receptor binding and transcriptional activation (90, 174). Taken in the context of heteromeric 105 PRMT2 complexes, it will be useful to see how different PRMT2 truncations may affect complex assemblies with nuclear receptors and other PRMTs. 4.4.4 Requirements for PRMT1/2 complexation Our previous work established that PRMT1 and PRMT2 can interact both in vitro and in cells (19); here we wanted to establish the molecular underpinnings responsible for this interaction. Using DSF, native gel electrophoresis, and enzymatic activity assays, we demonstrated evidence of a protein-protein interaction between PRMT1 and wildtype PRMT2. Our DSF data and native gel electrophoresis indicate that SAH and H3 peptide are required for PRMT1 and PRMT2 to interact. The most significant evidence for a PRMT1/2 protein-protein interaction is that PRMT2 influences PRMT1 catalytic activity towards specific histone proteins in a concentration-dependent manner. PRMT2 mutants revealed that PRMT2 SAM-binding and SH3 domains play a role in mediating interactions between PRMT1, PRMT2, and histone protein substrates. Further mechanistic studies should be performed to map out binding interactions within this enzyme-substrate complex. 4.4.5 PRMT2 impact on substrate specificity Previously, our group demonstrated that glutathione S-transferase (GST) fusion proteins of PRMT1 and PRMT2 form a complex that can augment PRMT1 activity towards histone H4 (19). We sought to further expand this observation by pursuing how PRMT2 influences PRMT1 methylation of core histones either individually, in complex as calf thymus histones, recombinant human histone octamers, or in unmodified mononucleosomes. Zheng and coworkers demonstrated that individual PRMT enzymes have differing substrate specificity towards free histones, histone octamers and nucleosomes (263). While individual PRMTs were unable to methylate recombinant unmodified mononucleosomes in vitro in their study, Vedadi and coworkers recently showed that H4K20 monomethylation in nucleosomes facilitated in vitro H4R3 methylation by PRMT1 (265). We also encountered differences in methylation depending on the macromolecular state of histones. Purified calf thymus histones H3 and H4 have several ε-N-methyllysine and ε-N-acetyllysine modifications in the N-terminus of both histone proteins [H3K9(Ac), H3K14(Ac), H3K23(Ac), H3K27(Me), H4K16(Ac), H4K20(Me)], but not histone H2A 106 (266â268). These modifications do not exist on the recombinantly expressed histones, histone octamers, or nucleosomes, and could be a major contributing factor to the discrepancies in observed methylation levels for PRMT1/2 (3, 106). Additionally, arginine methylation may occur prior to histone oligomerization or nucleosome formation, which could also explain why we see methylation pattern differences amongst the substrates tested. We speculate that further epigenetic modifications could further enhance the activity of the PRMT1/2 complex towards mononucleosomes both in vitro and in vivo. Nevertheless, we showed that combining these two PRMTs resulted in altered substrate specificity for PRMT1. 4.4.6 PRMT2 and the histone code PRMT2 is capable of augmenting PRMT1 and PRMT6 activity towards histone H2A depending on the context in which the substrate is presented. We showed that histone H2AR3 is the primary site for PRMT1/2 (Figure 4.5E). Another site for PRMT1 is H2AR11 (91, 260), whose methylation appeared to be modestly reduced by the H2AR11K mutation (Figure 4.5E). Histone H2A sites for PRMT6 are reported as H2AR3 and H2AR29 (91, 107). Since H2AR29me2a is enriched at genes repressed by PRMT6, we speculate that increased PRMT2 expression in cells may increase PRMT6-specific gene repression via increased H2AR29 methylation by a PRMT2/6 complex (260, 269). We demonstrate through DSF that PRMT2 is capable of binding the H2A peptide, in a manner we hypothesize is similar to how Mona Gads SH3 binds the SLP-76 peptide (Figure A31) (193). Given our results herein, it is possible that PRMT2 can modulate PRMT1 and PRMT6 activities in vivo towards H2A, resulting in substrate hypermethylation driving oncogenic expression. Further characterization is required to unravel the link between PRMT2 expression and H2A methylation in vivo. In addition to histone H2A methylation, we provide in vitro evidence for PRMT2 altering PRMT1âs ability to methylate histone H3 (Figure 4.5B). PRMT2 has been implicated either directly or indirectly with H3R8 methylation (184â186). It is possible that there may be conditions that enhance the catalytic activity of PRMT2 such that it can methylate H3R8 in a biologically relevant timeframe, but these conditions have yet to be identified and replicated in vitro. Our results provide an alternative 107 mechanism for H3R8 methylation as being deposited by a more active PRMT enzyme (e.g., PRMT1, PRMT6, or CARM1) whose activity is redirected by PRMT2 (19, 153, 154, 270). Beyond the core histones, the combination of PRMT2/CARM1 methylated a band in calf thymus histones consistent with the gel migration of linker histone H1 (Figure 4.6B); however, we were unable to methylate histone H1 as a recombinant solo substrate (data not shown). Histone H1 arginine methylation has yet to be reported. Of specific interest is the site H1R51, which can be citrullinated to regulate pluripotent activation (271). The methylation of this arginine could theoretically result in inhibition of citrullination and in turn inhibit cell pluripotency. It will be important to show that histone H1 is methylated in cells before there is any further speculation. 4.4.7 Concluding remarks In this study, we demonstrated the influence of PRMT2 in redirecting PRMT1 substrate specificity towards histone H2A. These findings suggest that direct arginine methylation may not be PRMT2âs primary function, but instead it may serve as an adaptor protein designed to mediate protein-protein interactions and recruit more active PRMTs for targeted substrate methylation. The PRMT2-dependent epigenetic modifications of histone proteins appear to rely on their macromolecular organization and quite possibly their modification status. Delving deeper into heteromeric PRMT2 complex activities will be imperative to understanding their biological implications and involvement with diseases like GBM, HCC, and RCC. 108 Chapter 5. Conclusions and future directions 5.1 Major thesis findings, significance, and impact The purpose of this research was to broaden the applicability of kinetic and thermodynamic assays used to study PRMTs with specific emphasis on PRMT2, its ligands, and protein interaction partners. In Chapter 2, we have provided a literature review of kinetic techniques used to study PRMT enzymes that addresses the need for more economical and environmentally friendly approaches. To address this need, I developed the filter binding and phosphor screening (FBAPS) assay to reduce the costs of consumables associated with performing radiometric PRMT assays while yielding comparable kinetic and inhibitory values. In Chapter 3, I evaluated the applicability of DSF towards investigating human PRMT2 thermal stability. Using DSF assays, I demonstrated how PRMT2 responds to thermal stress and explored how solution additives and small molecule inhibitors can influence PRMT2 thermal stability. In Chapter 4, I applied the techniques characterized in Chapters 2 and 3 towards better understanding how PRMT2 interacts with other proteins, including histones and other PRMT enzymes. 5.1.1 FBAPS strengths and limitations Unlike traditional radiometric filter binding assay, FBAPS replaces liquid scintillation counting with phosphor imaging, thus eliminating the use of single use plastic cuvettes and organic solvents. FBAPS affords the same kinetic parameters as the filter binding assay and can be quantified easily using low volumes of 14C-SAM spotted standards. Additionally, FBAPS moderately improves the throughput of the filter binding assay since the filter paper can be placed in a dot-blot apparatus to spot 96 unique conditions without needing to cut out pieces of filter paper. A limitation with FBAPS is the need for imaging instrumentation capable of phosphor imaging. Ultimately, FBAPS is a significant improvement of the filter binding assay that can allow researchers to evaluate PRMT kinetics and inhibition values while minimizing the usage of consumables. During our investigations, we identified that both FBAPS and the filter binding assay cannot accurately measure the activity of low activity methyltransferases due to radiolabeled SAM degradation products accumulating over long incubations at elevated temperature. Both radiolabeled SAM and SAM 109 degradation products can react with nucleophilic residues on protein substrates in a non-specific, uncatalyzed fashion, which cannot be distinguished from methylarginine by filter binding methods. Due to SAM degradation and non-specific SAM reactivity, we were not able to obtain any useful information about PRMT2 kinetics using the FBAPS assay. To separate methylarginine from other methylated amino acids generated by non-specific reactions with SAM, methylated protein substrates (with non-isotopically labelled methyl groups) can be acid hydrolyzed and analyzed by UPLC-MS/MS to selectively detect and quantitate each methylarginine species (19, 22, 35). A limitation to UPLC-MS/MS analysis is the requirement of sensitive mass spectrometry instrumentation and extensive sample preparation. 5.1.2 PRMT2 binding assays The work presented in Chapter 3 is the first characterization of human PRMT2 response to thermal stress. Using DSF, we demonstrated that PRMT2 displays a biphasic melt curve, and each melt transition is sensitive to the concentration of enzyme added. A limitation of this study is that we could not determine conclusively whether the biphasic melt was attributed to the unique melts of two independently folded domains or different oligomeric combinations of PRMT2. We identified that polyols and mildly acidic pH (5) improve PRMT2 thermal stability, while arginine and extremely acidic pH destabilized and potentially denatured PRMT2. We also demonstrated that hexa-histidine tagged human PRMT2 is less active than previously tested PRMT2 constructs, and the presence of stabilizing additives did not improve this activity. These findings demonstrate that external factors are capable of perturbing PRMT2 thermal stability, but those tested were shown to not improve PRMT2 catalytic activity. Unlike the excipients we tested, the ligands we evaluated elicited either no or slight PRMT2 thermal stabilization. The findings demonstrate that SAM binding to PRMT2 elicited a thermally stabilizing response, as expected from a SAM-dependent methyltransferase, while SAH binding did not. A limitation with thermal shift assays is that not all binding events influence the thermal stability of a protein, and such interactions do not result in melting curve shifts. A significant finding from evaluating PRMT2 response to ligands was the lack of thermal stabilization by MS023. Previous studies have claimed MS023 directly inhibits PRMT2 activity in cells, but failed to provide direct binding evidence of 110 this interaction (185, 186). While we cannot disprove MS023 binding to PRMT2 with DSF, our results demonstrate the need for more biophysical evaluations to determine whether MS023 does interact with and inhibit the activity of PRMT2. 5.1.3 PRMT2 protein-protein interactions In Chapter 4, we show a direct interaction between PRMT2 and three histone peptides (H2A, H3, H4) using DSF. We derived relative binding affinities for the interaction between PRMT2 and each histone peptide, showing a clear binding preference towards histone H3 peptide. While we failed to show thermal stabilization by SAH, we generated a mutation impairing PRMT2 ability to bind SAH/SAM (PRMT2H112Q) that abolished the thermal stabilization elicited by PRMT2 upon binding to the H3 peptide. Unlike the other PRMTs, we show that PRMT2 can bind protein substrates in the absence of cofactor, yet active site architecture responsible for cofactor binding is required for protein substrate binding. We additionally demonstrated the PRMT2 SH3 domain contributes to H3 peptide binding stabilization. These findings are essential to elucidation of the enzymatic mechanism of PRMT2 and suggest the SH3 domain plays a role in substrate binding, corroborating past findings showing PRMT2ÎSH3 had reduced activity (35). A limitation with this investigation is we do not provide a protein structure showing how PRMT2 might bind histone peptides. Previous work in the Frankel lab demonstrated that PRMT1 and PRMT2 interact both in vitro and in cells (19). We expanded on this observation by using DSF to characterize the protein-protein interactions between PRMT1 and PRMT2. Specifically, we showed that PRMT1 and PRMT2 experience a protein-protein interaction in a cofactor-, and SH3 domain-dependent manner. The ability to demonstrate protein-protein interactions using DSF has been demonstrated previously for proteins serving scaffolding roles in cells, but this is the first documented application of DSF towards interacting enzymes (261, 262). The ability to probe for protein-protein interactions by DSF allows researchers an alternative method to validate interactions and screen determinants for such interactions. Given the effect of PRMT2 on other PRMT kinetic capacities, it would be pertinent to further investigate protein-protein interactions between PRMT2 and other PRMTs. Additionally, DSF may also be used to evaluate heteromeric 111 interactions between PRMT2 and non-PRMT interacting partners, such as ER, AR, actin nucleator Cobl, and Sam68 to name a few (90, 174, 187, 200). DSF can be used to determine whether PRMT2 heteromeric interactions require the presence of additional ligands or specific buffer conditions. A limitation with using DSF to screen protein-protein interactions is that multiple experiments are required to gain quantitative information (binding affinity), instead of ITC where a single experiment can provide rich thermodynamic information. The disadvantage of ITC is necessity for expensive equipment and large quantities of protein. We corroborated our DSF findings of a PRMT1/2 protein-protein interactions with native PAGE and fluorescently tagged enzymes to demonstrate the proteins co-migrate in solution in a cofactor-dependent fashion. The major limitation with native-PAGE is it qualitatively assesses protein migration but is not proof of an interaction. While fluorescent fusions have been used before to investigate PRMT1/2 interactions in cells (via BiFC) (19), this is the first demonstration of a recombinantly expressed fluorescent PRMT2 fusion protein purified from bacteria. Previously, there were issues with expressing a functional fluorescent fusion of PRMT2 in vitro, but using Rosetta DE3 E. coli cells, I generated mCit-PRMT2 with an actively fluorescing fluorophore (likely due to the pRARE plasmid in RosettaDE3 that enhances the expression of eukaryotic proteins by supplying rare tRNA codons) (272). Having a fluorescently-tagged PRMT2 opens the door to multiple biophysical screening techniques, like FRET, fluorescence correlation spectroscopy, and microscale thermophoresis, representing a significant finding for the field (23, 273, 274). To further expand on the previous observation that PRMT2 can improve the catalytic activity of PRMT1 towards histone H4, we evaluated how different ratios of PRMT1 and PRMT2 can methylate calf thymus histones. We found that increasing amounts of PRMT2 promote the methylation of histone H4 and histone H2A, depending on the concentration of PRMT1 used, thus corroborating previous findings in our lab about histone H4 and demonstrating a novel methylation phenomenon towards histone H2A. Using PRMT2 site-directed mutagenesis, we show that the activation of PRMT1 by PRMT2 is dependent on the PRMT2 SH3 domain and PRMT2âs ability to bind SAM, but not PRMT2 catalytic activity. In fact, 112 we showed that the PRMT2 SH3 domain alone augmented the activity of PRMT1 towards histones H2A and H4. This is a significant finding that indicates that PRMT1 methylation of histones is impacted by presence of the PRMT2 N-terminus (containing the SH3 domain). Whether this observation extends to other SH3 domains is unknown. We further demonstrated that PRMT2 can influence the ability of PRMT3, PRMT4 (CARM1), PRMT5, PRMT6, PRMT7, and PRMT8 to methylate calf thymus histones. Taken together, these data demonstrate that PRMT2 plays a non-catalytic role in PRMT methylation of histones, potentially through its SH3 domain. To not limit our study to bovine histones, we also monitored how PRMT2 influences PRMT1 methylation of human histones, presented as individual recombinant histones, histone octamers, or nucleosomes. We demonstrated that PRMT2 reduces the KM of PRMT1 methylation of human recombinant histone H2A, H3, and H4, while increasing the Vmax of PRMT1 towards H2A and H3 when present in equimolar or 10-fold amounts (relative to PRMT1). Additionally, we found that PRMT2 negligibly impacts PRMT1 activity towards histone octamers, while dramatically impacting PRMT1 activity towards nucleosomes. This is the first demonstration of improved PRMT1 activity towards unmodified human nucleosomes in vitro and provides a significant step forward for PRMT2 enzymology. We observed higher activity of PRMT1/2 towards calf thymus histones, as opposed to the human histone octamers, indicating that pre-existing post translational modifications may make histones a better substrate for PRMT1/2 methylation. While H2A arginine methylation at R3 or R11 has not been implicated as either inhibitory or permissive of other epigenetic marks, H4R3me2a and H3R8me2a have both been implicated in a potential web of epigenetic modifications (14, 97, 153, 270). These observations could lead to a better understanding of how PRMTs influence the epigenetic code and can lead to the onset of disease. The greatest strength of this study is that multiple techniques were used to evaluate how PRMT2 interacts with both histones and other PRMT enzymes. While we demonstrate multiple novel findings that could explain how PRMT2 influences PRMT1 methylation of histones, this study did not conclusively demonstrate how all the proteins come together. To further probe how PRMT1 and PRMT2 interact with histones, structural biological techniques like cryogenic electron microscopy should be employed to 113 capture the protein complex and provide direct evidence of the interaction. Alternatively, site-directed mutagenesis could be employed to specifically mutate residues on PRMT2 that may be involved in either binding PRMT1 or histone proteins and evaluating their impact using techniques described in Chapter 4. While this study primarily focuses on the PRMT1 and PRMT2 interaction, PRMT2 has also been shown to interact with CARM1/PRMT4, PRMT6, and PRMT8 directly, providing multiple other proteins to characterize in vitro with PRMT2 to further understand PRMT2 complexation (39, 154). 5.2 Future directions 5.2.1 PRMT2 inhibitor design The findings in Chapters 3 and 4 provide important insights into molecules that can bind to and thermally stabilize PRMT2. Of all ligands tested, histone peptides, SAM, and sinefungin could elicit a thermally stabilizing response upon PRMT2. We did not experiment with PRMT2 in the presence of SAM and H3 peptide for two reasons. First, SAM is unstable and readily degrades in solution to produce several byproducts, including SAH that does not elicit a thermally stabilizing response upon PRMT2 and competes with SAM at the SAM-binding site. Second, PRMT2 can methylate full-length histone H3 protein, so in the presence of SAM and H3 peptide, PRMT2 could catalyze the formation of H3R8me2a peptide and SAH, leading to heterogeneity in ligands where it will be impossible to determine which combination of molecules is stabilizing PRMT2. To overcome this issue and to push forward in the development of inhibitors against PRMT2, bisubstrate or transition state analogues should be evaluated. Bisubstrate analogues are small molecules that mimic chemical features of two substrates that bind to an enzyme. PRMT bisubstrate inhibitors are designed to resemble SAM/SAH and an arginine residue. In the PRMT2 study conducted by Cavarelli and coworkers, they identified a synthetic molecule, Cp1, that consists of an adenosine scaffold connected to a guanidine with an ethylene spacer, acting as a PRMT bisubstrate analogue (141). Cp1 was demonstrated to inhibit murine PRMT2 and elicit a significantly greater thermal stabilization upon murine PRMT2 compared to SAH or sinefungin alone (141). It would be of paramount interest to evaluate the activity and thermal stabilization of human PRMT2 in presence of Cp1, especially given the discrepancy in responsiveness between murine and 114 human PRMT2 towards SAH. Cp1 is a generic PRMT bisubstrate analogue given it contains features that would be recognized widely amongst all PRMTs and would not yield the specificity required to selectively target PRMT2. Several bisubstrate inhibitors have been developed against PRMTs that have revealed important mechanistic features of PRMT binding specificity (26, 129, 138, 275, 276). DSF could be used to determine whether bisubstrate analogues interact with and stabilize PRMT2. Derivation of the DSF-derived KD of PRMT2 with the bisubstrate analogues could provide some indication of the affinity of interaction between each of the molecules with PRMT2 to determine which molecule binds with the highest affinity. Other groups have found success in the development of specific PRMT inhibitors through development of transition state analogues (134, 135). Transition state analogues are small molecules with chemical features that mimic the transition state of an enzyme-catalyzed chemical reaction. PRMT enzymes catalyze methyl transfer from SAM to arginine residues through a SN2 nucleophilic reaction, proceeding through formation of a pentavalent trigonal bipyramidal transition state between the methyl sulphonium of SAM and the terminal nitrogen on a guanidine ring of arginine residues (136). Martin and coworkers have shown through a variety of chemical biology studies that it is possible to develop specific PRMT transition state analogues using peptidomimetics (134, 277). The chemical scheme involved generating peptides of canonical substrates for the intended PRMT with an extra chemical appendage to the target arginine residue, such as H421R3 for PRMT1, HIV-Tat148-60R52 for PRMT1 and PRMT6, or PABP1447-459R454 and PABP1456-466R460 for CARM1 (134, 277). Martin and coworkers found that attaching an adenosine moiety with an ethylene linker to the terminal guanidine nitrogen of the arginine residues in these peptides serve as transition state analogues and bind with moderate affinity to their intended PRMT. Based on the work completed in Chapter 4 of this thesis, I identified a moderate affinity interaction between the histone H3 peptide and PRMT2. A future direction for the development of PRMT2 inhibitors could include synthesizing H3 peptide-based transition state analogues against PRMT2 and screening them through DSF and even the influencer assay to evaluate whether they compromise PRMT2 influence on PRMT1. Another group recently provided some evidence that PRMT2 was capable of methylating HIV-Tat R52 and R53, 115 suggesting the Tat peptidomimetics Martin and coworkers designed may also bind and inhibit PRMT2 activity (155, 277). 5.2.2 Interrogating the PRMT2 SH3 domain In Chapter 4, we demonstrate the PRMT2 SH3 domain may serve non-canonical roles in binding histones and activating PRMT1 methylation of histones. Other studies have demonstrated the PRMT2 SH3 domain can bind the proline rich stretches of PRMT8, BRD4, and the CobI actin nucleator (39, 187). In Chapter 4, we noted an acidic patch present on both the Mona/Gads SH3 domain and the PRMT2 SH3 domain located in the peptide binding groove that contact basic residues of SLP-76 (Figure A31C). We further identified a commonly shared motif between the SLP-76 peptide and histone peptides of: K/RxxK/R. A closer examination of the SH3 domains bound to SLP-76 showed a conserved glutamic acid in proximity with a lysine on SLP-76 to form a salt bridge. Three residues away from the lysine residue on the SLP-76 peptide is an arginine residue that interacts with either a leucine or threonine residue on the Mona/Gads or PRMT2 SH3 domain, respectively. To determine whether the identified residues on the PRMT2 SH3 domain can bind histone peptides, a series of PRMT2 site-directed mutants could be generated (using in silico modelling) and their DSF-binding affinities against the histone peptides measured. If a mutation is generated that promotes non-covalent interactions between the SH3 domain and the histone peptide, a smaller DSF-derived KD would be expected, whereas mutations that remove non-covalent interactions would increase the DSF-derived KD. Alternatively, a more direct approach to determining how PRMT2 binds histone peptides would be to use X-ray crystallography to solve the structure of human PRMT2 in complex with the histone peptide. If the histone peptides do bind the PRMT2 SH3 domain, they may serve as a crystallization aid that could yield a complete experimentally-derived crystal structure of PRMT2 with the SH3 domain and methyltransferase domain (278, 279). Currently, the complete molecular structure of dimeric human PRMT2 generated using Alpha Fold indicates there is poor confidence for positioning of the linker between the SH3 domain and the methyltransferase domain (Figure A32). Solving the complete structure of PRMT2 experimentally would provide direct evidence for how the SH3 domain is interacting with the methyltransferase domain. 116 Given the PRMT2 N-terminus could augment PRMT1 methylation of histone proteins, another direction to explore would be whether other SH3 domains are capable of binding histone peptides or activating PRMT methylation of histone proteins. This also begs the question if SH3 domains can improve the catalytic activity of other enzymes in cells. The findings in this thesis suggest that SH3 domains are involved in more than binding proline-rich motifs and their roles in biological systems require reconsideration and re-evaluations. 5.2.3 PRMT1/2 structural characterization There is a growing field of evidence demonstrating PRMT-PRMT heteromeric interactions play essential roles in cellular signaling and epigenetic regulation (19, 23, 153, 154). The exact mechanism of how PRMTs interact with one another remains enigmatic as there is no structural evidence of complex formation. Structural determination of a heteromeric PRMT1/2 complex using structural biology techniques, such as X-ray crystallography, NMR, or cryogenic electron microscopy (cryo-EM), would provide conclusive evidence that PRMTs do not exist solely as homomeric complexes. These techniques require that the proteins interact to form stable and homogenous complexes composed of both enzymes, that could be evaluated and optimized prior to structural determination. Previous studies have demonstrated that PRMT1 and PRMT2 can interact in vitro via co-IP, but this technique only demonstrates that the proteins can interact and does not demonstrate whether the proteins form stable and homogenous particles (19). Dynamic light scattering (DLS), and negative stain-EM can be used to screen whether PRMT1 and PRMT2 form stable complexes and the degree of homogeneity (280, 281). Particle homogeneity and stability can be optimized by addition of solution additives, such as a gradient fixation (GraFix) (282). Once conditions are identified that yield stable and homogenous PRMT1/2 complexes sample homogeneity (without disrupting the interaction between PRMT1/2 as measured by co-IP), the complexes can be analyzed by the structural biology technique of choice (X-ray crystallography, NMR, cryo-EM) (280). Solving the heteromeric structure of PRMT1/2 is a promising future direction that provide structural details for how these enzymes interact, potentially leading to insights into treatment of PRMT2-associated diseases. 117 5.2.4 PRMT1/2 and epigenetic crosstalk PRMT2 improves the activity of PRMT1 towards H2AR3, H3R8, and H4R3 when presented as recombinant proteins in solution. It was noted that PRMT1/2 demonstrated higher activity towards histone proteins purified from calf thymus (as octamers), as opposed to unmodified human histones. The N-termini of histones H2A, H3, and H4 are identical between bovine and human histones, indicating differences in arginine methylation by PRMT1/2 are not due to differences in sequence (283). This indicates the difference in methylation state between calf thymus and human histones can be attributed to pre-existing modifications on calf thymus histones (268). This suggests that the presence of pre-existing histone modifications may be pre-requisites for histone arginine methylation by PRMT1/2. Vedadi and coworkers demonstrated that PRMT1 had poor catalytic activity towards unmodified human nucleosomes but showed robust PRMT1 activity towards H4K20me1 methylated human nucleosomes (265). Numerous studies have been conducted to assess how post-translational modifications of histone proteins or peptides can impact the activity of epigenetic writers, demonstrating crosstalk between epigenetic modifications. Crosstalk between epigenetic marks can provide essential information about modifications that control chromatin organization and ultimately transcriptional access (284). Not only does crosstalk unveil important information about transcriptional states, but it can also indicate binding preferences for epigenetic writers, like PRMT1/2. To further expand on the substrate specificity of PRMT1/2, we propose screening the activity of this protein complex against a panel of differentially modified nucleosomes (with single modifications). EpiCypher provides the dCypherâ¢ âNucleosome Full Panelâ containing, unmodified control nucleosomes, nucleosomes connected with or without linker DNA, nucleosomes with variant histones, nucleosomes with oncogenic amino acid substitutions, nucleosomes with acidic patch mutants, tailless nucleosomes, and nucleosomes with defined single or combinatorial PTMs (285, 286). These nucleosomes could be screened against the PRMT1/2 complex by means of a high-throughput assay (like MTase-Glo methyltransferase assay kit from Promega) to identify initial hits and can be subsequently analyzed by gel radiography to visualize which histone is methylated. The results from such a screen 118 could identify histone modifications that promote or inhibit PRMT2-mediated histone methylation. From here, it will be possible to correlate PRMT2 activity with other histone marks. Identification of crosstalk between PRMT2-associated histone marks and other histone marks may provide insights into novel epigenetic signaling networks. 5.3 Conclusions PRMT2 is involved in numerous biological pathways and its overexpression has been associated with the onset or severity of several pathologies. Our characterization of PRMT2 thermal stability by DSF provides invaluable insights into which PRMT2 residues are involved in non-covalent interactions with ligand binding. 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Abbreviations are as follows: DDAH2 - dimethylarginine dimethylaminohydrolase 2; ESR1 â estrogen receptor 1; HNRNPA1 - heterogeneous nuclear ribonucleoprotein A1; HNRNPDL - heterogeneous nuclear ribonucleoprotein D-like; HNRNPUL1 - heterogeneous nuclear ribonucleoprotein U-like 1; L3MBTL3 â lethal (3) malignant brain tumor-like brain protein 3; PSRC1 â proline and serine rich coiled-coil 1; SETD1B - SET domain containing 1B; S100B â S100 calcium binding protein B. 153 Figure A3. PRMT2 GeneMANIA protein-protein interaction map. Edges connecting the nodes are color coordinated based on how the confidence of the interaction and evidence of the interaction. Abbreviations are as follows: AKR1B1, aldo-keto reductase family 1 member B; DPYSL3, dihydropyrimidinase-like 3; ESR2, estrogen receptor 2; FYN, FYN proto-oncogene, Src family tyrosine kinase; HNRNPUL1, heterogeneous nuclear ribonucleoprotein U-like 1; HTT, Huntingtin; GNB3, guanine nucleotide-binding protein 3; LTK, leukocyte receptor tyrosine kinase; METTL5, methyltransferase 5, N6-adenosine; NCOA6, nuclear receptor coactivator 6; NFKBIA, nuclear factor kappa B inhibitor alpha; N6AMT, N-6 adenine-specific DNA methyltransferase 1; PGR, progesterone receptor; PRRC2A, proline-rich coiled-coil 2A; RB1, retinoblastoma protein; SCARB1, scavenger receptor class B member 1; SHANK3, SH3 and multiple ankyrin repeat domains 3; TBCB, tubulin-folding cofactor B; THRB, thyroid hormone receptor beta. 154 Figure A4. Linear range determination using the FBAPS assay. Comparison of linear range of PRMT1 methylation of GST-GAR (A), and CARM1 methylation of GST-PABP1(437-488) (B). Enzyme-only reactions (black squares) and enzyme-catalyzed substrate reactions (red circles) are shown. Error bars represent standard deviation (n = 4). Figure A5. Optimizing total number of washes required for the filter paper used for FBAPS assay. Following application of the reactions onto P81 filter paper clamped in a 96-well dot-blot apparatus (as described in Figure 2.1), the P81 filter paper was dried at 37 ºC for 10 mins and then washed with 25 mL 50 mM NaCO3/NaHCO3 pH 9.0 buffer in a square dish for 5 mins. 1 mL of the wash buffer was mixed with 1 mL of Scintiverseâ¢ BD Cocktail in a scintillation vial and total disintegrations per minute (DPMs) were measured by scintillation counting. The liquid from each wash was disposed before the next wash. Error bars represent standard deviation (n = 2). The dotted line marks 100 DPM which is the UBC standard level of radiation deemed acceptable for cleanliness for wipe-tests. 0 1 2 3 4 5101102103104105# WashesDPM155 Figure A6. Evaluating the effects of increasing protein load on radioactive detection. (A) Loading key for spotting methylation reactions (n = 4) into 96-well dot-blot apparatus in which amount in µg GST-GAR of is indicated. (B) Coomassie blue-stained P81 filter paper spotted with methylation reactions containing PRMT1 (1.0 µM) and GST-GAR (11.5 µM) after 1 hr incubation with 50 µM 14C-SAM. Reactions were heat inactivated for 1 h at 70ËC prior to mixing with the amount of GST-GAR indicated in the loading key and spotted on the P81 filter paper. The initial amounts before additional GST-GAR were added are 0.81 µg PRMT1 and 2.25 µg GST-GAR. (C) Phosphor image obtained after exposing the Coomassie blue-stained filter paper (shown in (B)) to a storage phosphor screen for 1 h. 156 Figure A7. Evaluating the retention of methylated protein on P81 filter binding paper. (A) Schematic displaying how this assay was designed with stacked P81 filter paper. (B) Coomassie blue-stained P81 filter paper spotted with methylation reactions containing PRMT1 (1.0 µM) and GST-GAR (11.5 µM) after 1 hr incubation with 50 µM 14C-SAM. (C) Phosphor image obtained after exposing the Coomassie stained filter paper (shown in (B)) to a storage phosphor screen for 1 h. 157 Figure A8. PRMT2 purification analysis. (A) Chromatogram showing the A280 UV-Vis trace of PRMT2 elution from Ni-NTA HisTrap with 400 mM imidazole (green line indicating volume where buffer is switched from wash to elution buffer). Injection volume was clarified lysate from E. coli Arctic Express (DE3) cells overexpressing PRMT2. Ni-NTA fractions 5-7 were pooled for further purification via SEC. (B) Chromatogram showing the A280 UV-Vis trace of PRMT2 elution from HiLoad26/600 Superdex 200 pg SEC column. Gradations below the UV-Vis trace for both chromatograms indicate the volumes collected as fractions. SEC fractions 3-6 were pooled, concentrated, and buffer exchanged for kinetic and thermodynamic characterization (Chapters 2-4). (C) 10% SDS-PAGE analysis of the PRMT2 purification from E. coli Arctic Express by Ni-NTA and SEC chromatography. Fractions evaluated are the same as those monitored by the chromatograms in A and B. 158 CLUSTAL multiple sequence alignment by MUSCLE (3.8) NP_001526.2protein arginine N-methyltransferase 2 isoform 1 [Homo sapiens] AAI22564.1 protein arginine N-methyltransferase 2 isoform 1 [Mus musculus] NP_001526.2 MATSGDCPRSESQ-----GEEPAE--CSEAGLLQEG------VQPEEFVAIADYAATDET AAI22564.1 MEAPGEGPCSESQVIPVLEEDPVDYGC-EMQLLQDGAQLQLQLQPEEFVAIADYTATDET Human SH3 Domain NP_001526.2 QLSFLRGEKILILRQTTADWWWGERAGCCGYIPANHVGKHVDEYDPEDTWQDEEYFGSYG AAI22564.1 QLSFLRGEKILILRQTTADWWWGERAGCCGYIPANHLGKQLEEYDPEDTWQDEEYFDSYG Human Cofactor Binding Domain NP_001526.2 TLKLHLEMLADQPRTTKYHSVILQNKESLTDKVILDVGCGTGIISLFCAHYARPRAVYAV AAI22564.1 TLKLHLEMLADQPRTTKYHSVILQNKESLKDKVILDVGCGTGIISLFCAHHARPKAVYAV NP_001526.2 EASEMAQHTGQLVLQNGFADIITVYQQKVEDVVLPEKVDVLVSEWMGTCLLFEFMIESIL AAI22564.1 EASDMAQHTSQLVLQNGFADTITVFQQKVEDVVLPEKVDVLVSEWMGTCLLFEFMIESIL Human Beta Barrel Human Dimer arm NP_001526.2 YARDAWLKEDGVIWPTMAALHLVPCSADKDYRSKVLFWDNAYEFNLSALKSLAVKEFFSK AAI22564.1 YARDTWLKGDGIIWPTTAALHLVPCSAEKDYHSKVLFWDNAYEFNLSALKSLAIKEFFSR Human Beta Barrel NP_001526.2 PKYNHILKPEDCLSEPCTILQLDMRTVQISDLETLRGELRFDIRKAGTLHGFTAWFSVHF AAI22564.1 PKSNHILKPEDCLSEPCTILQLDMRTVQVPDLETMRGELRFDIQKAGTLHGFTAWFSVYF Human Beta Barrel NP_001526.2 QSLQEGQPPQVLSTGPFHP------------------------------TTHWKQTLFMM AAI22564.1 QSLEEGQPQQVLSTGPLHPFLGRTGCQCRGPRWSCQMRPVGDRMLLSCSTTHWKQTLFMM Human Beta Barrel NP_001526.2 DDPVPVHTGDVVTGSVVLQRNPVWRRHMSVALSWAVTSRQDPTSQKVGEKVFPIWR AAI22564.1 DDPVPVHTGDVVTGSVVLQRNPVWRRHMSVSLSWVVTSALDPTSQRVGEKVFPIWR Figure A9. CLUSTAL multiple PRMT sequence alignment by MUSCLE. PRMT2 from Homo sapiens, and Mus musculus were aligned and notable human domains were highlighted with structural PRMT feature above (in the same colour). Amino acids in M. musculus that do not align and have a different biophysical property compared to the H. sapiens sequence were bolded (within the catalytically relevant domains only). 159 Figure A10. PRMT2 methylation of GST-GAR. PRMT2 (1 µM) methylation reactions using 50 or 100 µM 14C-SAM and 0.45 mg/mL GST-GAR (16 h at 37 ºC). (A) GST-GAR only control, (B) PRMT2 only control, (C) PRMT2 and GST-GAR. Reactions were separated on a 10% SDS-PAGE and visualized by Coomassie staining (left), followed by phosphor imaging analysis after 2-week exposure (right). 160 Figure A11. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of osmolytes. Raw fluorescence from 7.5x SYPROTM Orange binding to (A) Increasing amounts of PRMT2 (5 â 15 µM) over a broad temperature range, or 5 µM PRMT2 in presence of increasing amounts of (B) glycerol (1 - 3%), (C) trehalose (0 â 400 mM), (D) glucose (0 â 400 mM) or (E) arginine (0 â 400 mM) (n = 3). Ligand-only controls included in each panel (n = 2). Tabulated melting temperatures and statistical analysis available in Tables A1 - 9. 161 Figure A12. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of pH. Raw fluorescence from 7.5x SYPROTM Orange binding to 10 µM PRMT2 in presence of (A) neutral to mildly acidic pH (5-8) or (B) mildly to highly acidic pH (2-5) (n = 3). Ligand-only controls included in each panel (n = 2). Tabulated melting temperatures and statistical analysis available in Appendix Tables 10 - 12. 162 Figure A13. Raw fluorescence of PRMT2 and SYPROTM Orange as a function of small molecule probes. Raw fluorescence from 7.5x SYPROTM Orange binding to 10 µM PRMT2 in presence of (A) SAH, (B) SAM, (C) Sinefungin, (D) Diamine 12, and (E) MS023 (+/- SAH) all at 500 µM (n = 3). Ligand-only controls included in each panel (n = 2). Tabulated melting temperatures and statistical analysis available in Appendix Tables 12 - 15. 163 Figure A14. Purity of recombinant PRMT enzymes used in Chapter 4. Enzymes used in Chapter 4 separated on 10% SDS-PAGE and stained by Coomassie Brilliant Blue. (A) Purity of non-fluorescently tagged PRMT variants used. (B) SDS-PAGE gel image from size-exclusion purification of fluorescently tagged PRMT variants used. Fractions 3 and 4 were pooled, concentrated, and buffer exchanged for mCer-PRMT1 and only fraction 5 was concentrated and buffer-exchanged for mCit-PRMT2. 164 Figure A15. Impact of ligands on PRMT2 thermal stability. Raw fluorescence from 7.5x SYPROTM Orange binding to PRMT2 (5 µM) over a broad temperature range in presence of increasing concentrations of (A) SAH (purple), (B) H2A peptide (blue), (C) H3 peptide (orange), or (D) H4 peptide (green) (n = 3). Peptide-only controls are shown in (E). Ligand-only controls included in each figure (n = 2). Tabulated melting temperatures and statistical analysis available in Tables A16 and A18. 165 Figure A16. PRMT2 bisubstrate thermal stabilization analysis. PRMT2 (5 µM) was incubated with saturating amounts (500 µM) of SAH, and/or H3 peptide and the thermal stabilization of these ligands on PRMT2 was assessed by DSF. First-derivative melting traces (in relative florescence units per second (RFU/s)) are the average of three replicates; error bars show standard deviations. Tabulated melting temperatures and statistical analysis available in Tables A18 and A19. Figure A17. Thermal melt of PRMT2H112Q in DSF. (A) Raw fluorescence from 7.5x SYPROTM Orange binding to 5 µM PRMT2H112Q over a broad temperature range in presence or absence of saturating H3 peptide and/or SAH (500 µM). (B) First-derivative melting curves (in relative florescence units per second (RFU/s)) of PRMT2H112Q melt curves shown in A. Tabulated melting temperatures and statistical analysis available in Tables A20 and A21. 166 Figure A18. Thermal melt of PRMT2ÎSH3 in DSF. (A) Raw fluorescence from 7.5x SYPROTM Orange binding to 5 µM PRMT2ÎSH3 over a broad temperature range in presence or absence of saturating H3 peptide and/or SAH (500 µM). (B) Normalized fluorescence of SYPROTM Orange and PRMT2ÎSH3 with increasing concentration of H3 peptide (0-500 µM). (C) First-derivative melting curves (in relative florescence units per second (RFU/s)) of PRMT2 ÎSH3 melt curves shown in A. (D) Normalized fluorescence of SYPROTM Orange and PRMT2ÎSH3 in the presence or absence of SAH and/or H3 peptide (n = 3). (E) DSF-derived KD for H3 peptide binding to PRMT2ÎSH3. Ligand-only controls included in (A). Tabulated melting temperatures and statistical analysis available in Tables A22 - 24. 167 Figure A19. Impact of H3 peptide on PRMT1 thermal stability. (A) Raw fluorescence and (B) first-derivative data (in relative florescence units per second (RFU/s)) from 7.5x SYPROTM Orange binding to 5 µM PRMT1 over a broad temperature range in the presence or absence of saturating H3 peptide and/or SAH (500 µM). (C) First-derivative data (in relative florescence units per second (RFU/s)) from 5 µM PRMT1 in presence of increasing H3 peptide while holding SAH constant (500 µM) and (D) DSF-derived KD for H3 peptide binding to PRMT1. Tabulated melting temperatures and statistical analysis available in Tables A25 - 27. 168 Figure A20. PRMT1/2 complex thermal stability. Raw fluorescence from 7.5x SYPROTM Orange binding to (A) 5 µM PRMT1 and 5 µM PRMT2 over a broad temperature range in absence of cofactor, (B) presence of SAH (500 µM), (C) H3 peptide (500 µM), or (D) both SAH and H3 peptide (500 µM) (n = 3). Ligand-only controls included in each panel (n = 2). Tabulated melting temperatures and statistical analysis available in Tables A28 and A29. 169 Figure A21. PRMT1/PRMT2ÎSH3 complex thermal stability. Raw fluorescence from 7.5x SYPROTM Orange binding to 5 µM PRMT1 and 5 µM PRMT2ÎSH3 over a broad temperature range (A) in absence of ligands, (B) 500 µM SAH, (C) 500 µM H3 peptide, or (D) both SAH and H3 peptide at 500 µM each (n = 3). Ligand-only controls included in each figure (n = 2). Tabulated melting temperatures and statistical analysis available in Tables A30 and A31. 170 Figure A22. PRMT1/PRMT2H112Q complex thermal stability. Raw fluorescence from 7.5x SYPROTM Orange binding to 5 µM PRMT1 and 5 µM PRMT2H112Q over a broad temperature range (A) in absence of ligands, (B) 500 µM SAH, (C) 500 µM H3 peptide, or (D) both SAH and H3 peptide at 500 µM each (n = 3). Ligand-only controls included in each figure (n = 2). Tabulated melting temperatures and statistical analysis available in Tables A32 and A33. 171 Figure A23. PRMT1/2 complex thermal stability varying protein concentrations. First-derivative of fluorescence with respect to temperature (in relative florescence units per second (RFU/s)) for 7.5x SYPROTM Orange binding to 1-10 µM PRMT1 and 1-10 µM PRMT2 over a broad temperature range in absence of ligands, 500 µM SAH, 500 µM H3 peptide, or both SAH and H3 peptide at 500 µM each as indicated (n = 3). Tabulated melting temperatures available in Table A34. 172 Figure A24. Impact of storage buffer on PRMT1 methylation of calf thymus histones. PRMT1 (1 µM) methylation reactions using 10 µM 14C-SAM and 10 µg calf thymus histones with increasing amounts of storage buffer (1x =100 mM HEPES-KOH pH 8.0, 200 mM NaCl, 1 mM DTT, 10% glycerol, 2 mM EDTA). Reactions were separated on a 16.5% Tricine SDS-PAGE and visualized by Coomassie staining (top), followed by phosphor imaging analysis after 24-h exposure (bottom). 173 Figure A25. PRMT1E153Q/2 methylation of calf thymus histones. PRMT1E153Q (1 µM) methylation reactions using 10 µM 14C-SAM and 10 µg calf thymus histones with increasing amounts of PRMT2. Reactions were separated on a 16.5% Tricine SDS-PAGE and visualized by Coomassie staining (top), followed by phosphor imaging analysis after 48-h exposure (bottom). 174 Figure A26. Gel radiography analysis of calf thymus histone methylation by PRMT1/2. PRMT1 (100 nM) was pre-incubated with PRMT2 (0, 100, 1000, 10000 nM) prior to reaction with 10 µM 14C-SAM and 0-300 µg/mL calf thymus histones for 2 hrs. (A) Reactions were separated on 16.5% Tricine-SDS-PAGE, Coomassie stained (top) dried down and exposed to storage phosphor for 12 h before imaging (bottom). Tabulated values from the densitometry analysis can be found in Table A35. 175 BLOT A 1 2 3 4 5 6 7 8 9 10 11 12 A H2A (25 µM) PRMT1/2 (1:0) PRMT1/2 (1:0) + 0.1 µM H2A PRMT1/2 (1:0) + 0.5 µM H2A B PRMT1/2 (1:0) + 1 µM H2A PRMT1/2 (1:0) + 5 µM H2A PRMT1/2 (1:0) + 10 µM H2A PRMT1/2 (1:0) + 15 µM H2A C PRMT1/2 (1:0) + 20 µM H2A PRMT1/2 (1:0) + 25 µM H2A PRMT1/2 (1:100) PRMT1/2 (1:100) + 0.1 µM H2A D PRMT1/2 (1:100) + 0.5 µM H2A PRMT1/2 (1:100) + 1 µM H2A PRMT1/2 (1:100) + 5 µM H2A PRMT1/2 (1:100) + 10 µM H2A E PRMT1/2 (1:100) + 15 µM H2A PRMT1/2 (1:100) + 20 µM H2A PRMT1/2 (1:100) + 25 µM H2A BLOT B 1 2 3 4 5 6 7 8 9 10 11 12 A H2A (25 µM) PRMT1/2 (1:1) PRMT1/2 (1:1) + 0.1 µM H2A PRMT1/2 (1:1) + 0.5 µM H2A B PRMT1/2 (1:1) + 1 µM H2A PRMT1/2 (1:1) + 5 µM H2A PRMT1/2 (1:1) + 10 µM H2A PRMT1/2 (1:1) + 15 µM H2A C PRMT1/2 (1:1) + 20 µM H2A PRMT1/2 (1:1) + 25 µM H2A PRMT1/2 (1:10) PRMT1/2 (1:10) + 0.1 µM H2A D PRMT1/2 (1:10) + 0.5 µM H2A PRMT1/2 (1:10) + 1 µM H2A PRMT1/2 (1:10) + 5 µM H2A PRMT1/2 (1:10) + 10 µM H2A E PRMT1/2 (1:10) + 15 µM H2A PRMT1/2 (1:10) + 20 µM H2A PRMT1/2 (1:10) + 25 µM H2A Figure A27. FBAPS analysis of H2A methylation by PRMT1/2. PRMT1 (100 nM) was pre-incubated with PRMT2 (0, 100, 1000, 10000 nM) prior to reaction with 10 µM 14C-SAM and the indicated concentration of recombinant H2A for 2 h. Reactions were spotted on P81 filter paper (n = 3) according to grid (top) via dot-blot apparatus under vacuum, washed 5x in 50 mM sodium phosphate pH 9.0, dried down and exposed to storage phosphor for 12 h before imaging (bottom). Tabulated densitometry data available in Table A36. 176 BLOT A 1 2 3 4 5 6 7 8 9 10 11 12 A H3 (25 µM) PRMT1/2 (1:0) PRMT1/2 (1:0) + 0.1 µM H3 PRMT1/2 (1:0) + 0.5 µM H3 B PRMT1/2 (1:0) + 1 µM H3 PRMT1/2 (1:0) + 5 µM H3 PRMT1/2 (1:0) + 10 µM H3 PRMT1/2 (1:0) + 15 µM H3 C PRMT1/2 (1:0) + 20 µM H3 PRMT1/2 (1:0) + 25 µM H3 PRMT1/2 (1:100) PRMT1/2 (1:100) + 0.1 µM H3 D PRMT1/2 (1:100) + 0.5 µM H3 PRMT1/2 (1:100) + 1 µM H3 PRMT1/2 (1:100) + 5 µM H3 PRMT1/2 (1:100) + 10 µM H3 E PRMT1/2 (1:100) + 15 µM H3 PRMT1/2 (1:100) + 20 µM H3 PRMT1/2 (1:100) + 25 µM H3 BLOT B 1 2 3 4 5 6 7 8 9 10 11 12 A H3 (25 µM) PRMT1/2 (1:1) PRMT1/2 (1:1) + 0.1 µM H3 PRMT1/2 (1:1) + 0.5 µM H3 B PRMT1/2 (1:1) + 1 µM H3 PRMT1/2 (1:1) + 5 µM H3 PRMT1/2 (1:1) + 10 µM H3 PRMT1/2 (1:1) + 15 µM H3 C PRMT1/2 (1:1) + 20 µM H3 PRMT1/2 (1:1) + 25 µM H3 PRMT1/2 (1:10) PRMT1/2 (1:10) + 0.1 µM H3 D PRMT1/2 (1:10) + 0.5 µM H3 PRMT1/2 (1:10) + 1 µM H3 PRMT1/2 (1:10) + 5 µM H3 PRMT1/2 (1:10) + 10 µM H3 E PRMT1/2 (1:10) + 15 µM H3 PRMT1/2 (1:10) + 20 µM H3 PRMT1/2 (1:10) + 25 µM H3 Figure A28. FBAPS analysis of H3 methylation by PRMT1/2. PRMT1 (100 nM) was pre-incubated with PRMT2 (0, 100, 1000, 10000 nM) prior to reaction with 10 µM 14C-SAM and the indicated concentration of recombinant H3 for 2 h. Reactions were spotted on P81 filter paper (n = 3) according to grid (top) via dot-blot apparatus under vacuum, washed 5x in 50 mM sodium phosphate pH 9.0, dried down and exposed to storage phosphor for 12 h before imaging (bottom). Tabulated densitometry data available in Table A37. 177 BLOT A 1 2 3 4 5 6 7 8 9 10 11 12 A H4 (20 µM) PRMT1/2 (1:0) PRMT1/2 (1:0) + 0.1 µM H4 PRMT1/2 (1:0) + 0.5 µM H4 B PRMT1/2 (1:0) + 1 µM H4 PRMT1/2 (1:0) + 5 µM H4 PRMT1/2 (1:0) + 10 µM H4 PRMT1/2 (1:0) + 15 µM H4 C PRMT1/2 (1:0) + 20 µM H4 PRMT1/2 (1:1) PRMT1/2 (1:1) + 0.1 µM H4 PRMT1/2 (1:1) + 0.5 µM H4 D PRMT1/2 (1:1) + 1 µM H4 PRMT1/2 (1:1) + 5 µM H4 PRMT1/2 (1:1) + 10 µM H4 PRMT1/2 (1:1) + 15 µM H4 E PRMT1/2 (1:1) + 20 µM H4 PRMT1/2 (1:10) PRMT1/2 (1:10) + 0.1 µM H4 PRMT1/2 (1:10) + 0.5 µM H4 F PRMT1/2 (1:10) + 1 µM H4 PRMT1/2 (1:10) + 5 µM H4 PRMT1/2 (1:10) + 10 µM H4 PRMT1/2 (1:10) + 15 µM H4 G PRMT1/2 (1:10) + 20 µM H4 PRMT1/2 (1:100) PRMT1/2 (1:100) + 0.1 µM H4 PRMT1/2 (1:100) + 0.5 µM H4 H PRMT1/2 (1:100) + 1 µM H4 PRMT1/2 (1:100) + 5 µM H4 PRMT1/2 (1:100) + 10 µM H4 PRMT1/2 (1:100) + 15 µM H4 BLOT B 1 2 3 4 5 6 7 8 9 10 11 12 A PRMT1/2 (1:100) + 20 µM H4 Figure A29. FBAPS analysis of H4 methylation by PRMT1/2. PRMT1 (100 nM) was pre-incubated with PRMT2 (0, 100, 1000, 10000 nM) prior to reaction with 10 µM 14C-SAM and the indicated concentration of recombinant H3 for 2 h. Reactions were spotted on P81 filter paper (n = 3) according to grid (top) via dot-blot apparatus under vacuum, washed 5x in 50 mM sodium phosphate pH 9.0, dried down and exposed to storage phosphor for 48 h before imaging (bottom). Tabulated densitometry data available in Table A38. 178 Figure A30. FBAPS quantitation of individual histone methylation by PRMT2. (A) PRMT2 (10 µM) reaction with 10 µM 14C-SAM and 25 µM of the indicated concentration of recombinant histone for 2 h. (B) The same blot shown in Supporting Figure 19 but now exposed with a 14C-SAM ladder to quantify absolute levels of C-14. Reactions were spotted on P81 filter paper (n = 3) according to grid (top) via dot-blot apparatus under vacuum, washed 5x in 50 mM sodium phosphate pH 9.0, dried down and exposed to storage phosphor for 72 h before imaging. Both filter papers exposed with same 14C-SAM ruler to show PRMT2 reaction with histones and 14C-SAM yields no detectable activity compared to when PRMT1 is present. *The same blot was used from Appendix Figure 28 for comparison purpose against the PRMT2 only controls to show the differences in intensity between a PRMT1/2 reaction against H4 and PRMT2 against H4 only* 179 Figure A31. Possible PRMT2 SH3 domain ligands. (A) The human PRMT2 SH3 domain and the Mona/Gads SH3C domain share 31% sequence identity and 47% sequence similarity (+). (B) The Mona/Gads SH3C domain (tan) (PDB 1OEB; amino acids 266-316) in complex with a peptide portion of the T-cell signal transducer SLP-76 (brown) with the R/KXXK motif highlighted (salmon) superimposed with the PRMT2 SH3 domain (PDB 1X2P; amino acids 33-83) showing an r.m.s.d. of 0.975 Ã were rendered using Chimera (UCSF). (C) The superimposition described in (B) with the PRMT2 SH3 domain electrostatic surface map is showing basic residues of the RXXK motif interacting with acidic surfaces. (D) The RXXK motif (bolded) found in the SLP-76 sequence can also be found as (R/K)XX(R/K) in bovine histone N-termini, suggesting that the PRMT2 SH3 domain may be able to interact with these sequences. Bovine histone sequences are shown since calf thymus histones are used as methylation substrates in our assays. 180 Figure A32. AlphaFold structure of dimeric human PRMT2. (A) Ribbon diagram of the dimeric structure of human PRMT2 generated using AlphaFold Colab2, colored according to chain identity on the left (monomer A, green; monomer B, cyan), or according to residue confidence score (pLDDT) on the right (see color scale in B). (B) Confidence in residue positioning for the model shown in (A) predicted by AlphaFold Colab2. AB181 Appendix Tables Table A1. Derived melting temperature (Tm) for PRMT2 from Figure 3.1C and D. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept corresponding to the local maxima observed on the first-derivative plot. R1-R3 are replicates, AVE is average, and SD is standard deviation. Statistical comparisons are shown in Table A2. 5 µM PRMT2 10 µM PRMT2 15 µM PRMT2 Tm1 (ºC) Tm1 (ºC) Tm1 (ºC) R1 R2 R3 R1 R2 R3 R1 R2 R3 46.63 46.34 46.29 48.63 48.45 48.89 54.50 54.22 54.65 AVE SD N AVE SD N AVE SD N 46.42 0.15 3.00 48.66 0.18 3.00 54.46 0.18 3.00 Tm2 (ºC) Tm2 (ºC) Tm2 (ºC) R1 R2 R3 R1 R2 R3 R1 R2 R3 49.98 49.92 49.56 52.05 51.53 51.62 58.45 58.45 58.17 AVE SD N AVE SD N AVE SD N 49.82 0.18 3.00 51.73 0.23 3.00 58.36 0.13 3.00 5 µM PRMT2 + 2% Glycerol 5 µM PRMT2 + 3% Glycerol Tm1 (ºC) Tm1 (ºC) R1 R2 R3 48.90 48.99 48.84 AVE SD N 48.91 0.06 3.00 Tm2 (ºC) Tm2 (ºC) R1 R2 R3 R1 R2 R3 52.06 52.11 51.79 53.91 53.82 54.50 AVE SD N AVE SD N 51.99 0.14 3.00 54.07 0.30 3.00 182 Table A2. One-way ANOVA statistical testing evaluating the melting temperature (Tm) from Figure 3.1C and D. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm Tm1 (ºC) Tm2 (ºC) Below threshold? Summary Adjusted P Value PRMT2 5 µM vs. PRMT2 10 µM 2.24 â Yes **** 2.49 x 10-6 PRMT2 5 µM vs. PRMT2 15 µM 8.04 â Yes **** 1.01 x 10-10 PRMT2 5 µM vs. PRMT2 5 µM + 2% Glycerol 2.49 â Yes **** 1.08 x 10-6 PRMT2 10 µM vs. PRMT2 5 µM 5.80 â Yes **** 1.36 x 10-9 PRMT2 10 µM vs. PRMT2 5 µM + 2% Glycerol 0.25 â No ns 0.79 PRMT2 15 µM vs. PRMT2 µM + 2% Glycerol -5.55 â Yes **** 1.94 x 10-9 PRMT2 5 µM vs. PRMT2 10 µM 1.91 â Yes **** 3.24 x 10-5 PRMT2 5 µM vs. PRMT2 15 µM 8.54 â Yes **** 1.68 x 10-11 PRMT2 5 µM vs. PRMT2 5 µM + Glycerol 2% 2.17 â Yes **** 1.04 x 10-5 PRMT2 5 µM vs. PRMT2 5 µM + Glycerol 3% 4.26 â Yes **** 1.63 x 10-8 PRMT2 10 µM vs. PRMT2 15 µM 6.62 â Yes **** 2.08 x 10-10 PRMT2 10 µM vs. PRMT2 5 µM + Glycerol 2% 0.25 â No ns >0.99 PRMT2 10 µM vs. PRMT2 5 µM + Glycerol 3% 2.34 â Yes **** 5.02 x 10-6 PRMT2 15 µM vs. PRMT2 5 µM + Glycerol 2% -6.37 â Yes **** 3.06 x 10-10 PRMT2 15 µM vs. PRMT2 5 µM + Glycerol 3% -4.28 â Yes **** 1.54 x 10-8 PRMT2 5 µM + Glycerol 2% vs. PRMT2 5 µM + Glycerol 3% 2.09 â Yes **** 1.45 x 10-5 183 Table A3. Derived melting temperature for PRMT2 (Tm1 and Tm2) from Figure 3.2A. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. [Trehalose] (mM) Tm1 (ºC) Tm2 (ºC) 0 R1 R2 R3 R1 R2 R3 46.07 46.06 46.58 49.85 50.75 49.69 Ave SD N Ave SD N 46.24 0.24 3 50.09 0.47 3 10 R1 R2 R3 R1 R2 R3 46.76 47.07 47.08 50.1 50.81 50.56 Ave SD N Ave SD N 46.97 0.15 3 50.49 0.29 3 50 R1 R2 R3 R1 R2 R3 47.77 47.79 47.64 51.01 51.47 50.97 Ave SD N Ave SD N 47.74 0.07 3 51.15 0.23 3 100 R1 R2 R3 R1 R2 R3 48.17 47.82 47.8 51.48 50.97 50.66 Ave SD N Ave SD N 47.93 0.17 3 51.04 0.34 3 200 R1 R2 R3 R1 R2 R3 49.44 48 49.08 51.71 52.01 Ave SD N Ave SD N 48.84 0.61 3 51.86 0.15 3 300 R1 R2 R3 R1 R2 R3 48.64 48.08 48.81 52.06 52 52.13 Ave SD N Ave SD N 48.51 0.31 3 52.06 0.05 3 400 R1 R2 R3 R1 R2 R3 49.99 50.04 49.83 54.43 54.29 53.6 Ave SD N Ave SD N 49.96 0.09 3 54.11 0.36 3 184 Table A4. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.2A. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Tm1 (°C) Tm2 (°C) Summary Adjusted P Value PRMT2 vs. PRMT2 + 10 mM Trehalose 0.73 â * 4.9 x 10-2 PRMT2 vs. PRMT2 + 50 mM Trehalose 1.50 â *** 1.12 x 10-4 PRMT2 vs. PRMT2 + 100 mM Trehalose 1.69 â **** 3.06 x 10-5 PRMT2 vs. PRMT2 + 200 mM Trehalose 2.60 â **** 1.77 x 10-7 PRMT2 vs. PRMT2 + 300 mM Trehalose 2.27 â **** 9.58 x 10-7 PRMT2 vs. PRMT2 + 400 mM Trehalose 3.72 â **** 1.68 x 10-9 PRMT2 vs. PRMT2 + 10 mM Trehalose 0.40 â ns 0.75 PRMT2 vs. PRMT2 + 50 mM Trehalose 1.06 â ** 4.12 x 10-3 PRMT2 vs. PRMT2 + 100 mM Trehalose 0.95 â ** 9.91 x 10-3 PRMT2 vs. PRMT2 + 200 mM Trehalose 1.77 â **** 2.59 x 10-5 PRMT2 vs. PRMT2 + 300 mM Trehalose 1.97 â **** 7.59 x 10-6 PRMT2 vs. PRMT2 + 400 mM Trehalose 4.02 â **** 9.05 x 10-10 185 Table A5. Derived melting temperature (Tm1 and Tm2) for PRMT2 from Figure 3.2B. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. [Glucose] (mM) Tm1 (ºC) Tm2 (ºC) 0 R1 R2 R3 R1 R2 R3 47.16 46.43 47.14 51.01 50.22 50.92 Ave SD N Ave SD N 46.91 0.34 3 50.72 0.35 3 10 R1 R2 R3 R1 R2 R3 47.71 47.28 46.93 51.34 50.73 50.54 Ave SD N Ave SD N 47.31 0.32 3 50.87 0.34 3 50 R1 R2 R3 R1 R2 R3 47.35 47.34 47.03 51.01 50.95 50.95 Ave SD N Ave SD N 47.24 0.15 3 50.97 0.03 3 100 R1 R2 R3 R1 R2 R3 47.34 47.29 47 51.17 50.99 50.86 Ave SD N Ave SD N 47.21 0.15 3 51.01 0.13 3 200 R1 R2 R3 R1 R2 R3 47.14 47.57 47.15 51.94 51.19 51.41 Ave SD N Ave SD N 47.29 0.2 3 51.51 0.31 3 300 R1 R2 R3 R1 R2 R3 48.31 48.47 48.14 52.25 52.51 52.07 Ave SD N Ave SD N 48.31 0.14 3 52.28 0.18 3 400 R1 R2 R3 R1 R2 R3 51.91 50.44 55.13 53.94 Ave SD N Ave SD N 51.17 0.74 2 54.54 0.6 2 186 Table A6. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.2B. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Tm1 (°C) Tm2 (°C) Summary Adjusted P Value PRMT2 vs. PRMT2 + 10 mM Glucose -0.40 â ns 0.78 PRMT2 vs. PRMT2 + 50 mM Glucose -0.33 â ns >0.99 PRMT2 vs. PRMT2 + 100 mM Glucose -0.30 â ns >0.99 PRMT2 vs. PRMT2 + 200 mM Glucose -0.38 â ns 0.89 PRMT2 vs. PRMT2 + 300 mM Glucose -1.40 â *** 4.67 x 10-4 PRMT2 vs. PRMT2 + 400 mM Glucose -4.26 â **** 5.96 x 10-9 PRMT2 vs. PRMT2 + 10 mM Glucose -0.15 â ns >0.99 PRMT2 vs. PRMT2 + 50 mM Glucose -0.25 â ns >0.99 PRMT2 vs. PRMT2 + 100 mM Glucose -0.29 â ns >0.99 PRMT2 vs. PRMT2 + 200 mM Glucose -0.79 â * 3.56 x 10-2 PRMT2 vs. PRMT2 + 300 mM Glucose -1.56 â *** 1.23 x 10-4 PRMT2 vs. PRMT2 + 400 mM Glucose -3.82 â **** 1.62 x 10-10 187 Table A7. Derived melting temperature for PRMT2 (Tm1 and Tm2) from Figure 3.2C. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. [Arginine] (mM) Tm1 (ºC) Tm2 (ºC) 0 R1 R2 R3 R1 R2 R3 47.08 47.84 47.12 50.79 50.99 50.570 Ave SD N Ave SD N 47.35 0.35 3 50.78 0.17 3 10 R1 R2 R3 R1 R2 R3 47.52 47.37 47.40 51.05 50.92 50.98 Ave SD N Ave SD N 47.43 0.06 3 50.98 0.05 3 100 R1 R2 R3 48.92 48.60 48.70 Ave SD N 48.74 0.14 3 200 R1 R2 R3 43.10 44.94 44.66 Ave SD N 44.23 0.81 3 Table A8. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure 3.2C. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2 vs. PRMT2 + 10 mM Arginine -0.08 ns >0.99 PRMT2 vs. PRMT2 + 100 mM Arginine -1.39 * 1.57 x 10-2 PRMT2 vs. PRMT2 + 200 mM Arginine 3.12 **** 8.20 x 10-2 Table A9. Unpaired studentâs statistical testing evaluating the melting temperatures (Tm2) from Figure 3.2C. A statistical level of significance cut-off <0.05 was used. Unpaired t test with Welch's correction P value 0.17 P value summary ns Significantly different (P < 0.05)? No One- or two-tailed P value? Two-tailed Welch-corrected t, df t=1.955, df=2.343 188 Table A10. Derived melting temperature for PRMT2 (Tm) from Figure 3.3A and B. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. pH Tm (ºC) 3 R1 R2 R3 46.54 45.15 45.68 Ave SD N 45.79 0.57 3.00 4 R1 R2 R3 62.47 62.27 62.48 Ave SD N 62.40 0.09 3.00 5 R1 R2 R3 62.96 63.26 63.26 Ave SD N 63.16 0.14 3.00 5.5 R1 R2 R3 60.61 60.92 60.97 Ave SD N 60.83 0.16 3.00 6 R1 R2 R3 60.32 60.25 59.47 Ave SD N 60.02 0.38 3.00 6.5 R1 R2 R3 56.13 55.72 55.84 Ave SD N 55.90 0.17 3.00 7 R1 R2 R3 55.15 55.10 55.35 Ave SD N 55.20 0.11 3.00 7.5 R1 R2 R3 53.41 53.34 53.24 Ave SD N 53.33 0.07 3.00 8 R1 R2 R3 53.17 53.59 52.88 Ave SD N 53.21 0.29 3.00 189 Table A11. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.3A and 3.3B. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2 pH 8 vs. PRMT2 pH 3 -7.42 **** <1.0 x 10-15 PRMT2 pH 8 vs. PRMT2 pH 4 9.19 **** <1.0 x 10-15 PRMT2 pH 8 vs. PRMT2 pH 5 9.95 **** <1.0 x 10-15 PRMT2 pH 8 vs. PRMT2 pH 5.5 7.62 **** <1.0 x 10-15 PRMT2 pH 8 vs. PRMT2 pH 6 6.81 **** <1.0 x 10-15 PRMT2 pH 8 vs. PRMT2 pH 6.5 2.69 **** 2.95 x 10-9 PRMT2 pH 8 vs. PRMT2 pH 7 1.99 **** 3.24 x 10-7 PRMT2 pH 8 vs. PRMT2 pH 7.5 0.12 ns >0.99 Table A12. Derived melting temperature for PRMT2 (Tm) from Figure 3.4B, C, D. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Treatment Tm1 (ºC) PRMT2 R1 R2 R3 53.59 53.38 53.31 AVE SD N 53.43 0.12 3.00 SAM R1 R2 R3 54.27 54.64 54.61 AVE SD N 54.51 0.17 3.00 Sinefungin R1 R2 R3 54.71 54.54 54.54 AVE SD N 54.59 0.08 3.00 Diamine 12 R1 R2 R3 53.63 53.81 53.57 AVE SD N 53.67 0.10 3.00 190 Table A13. Derived melting temperature for PRMT2 (Tm) from Figure 3.4 A, E. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Treatment Tm1 (ºC) PRMT2 R1 R2 R3 53.05 52.67 52.59 AVE SD N 52.77 0.20 3.00 SAH R1 R2 R3 53.09 53.09 53.20 AVE SD N 53.13 0.05 3.00 DMSO R1 R2 R3 53.25 53.22 53.34 AVE SD N 53.27 0.05 3.00 MS023 R1 R2 R3 53.33 53.04 53.05 AVE SD N 53.14 0.14 3.00 SAH + MS023 R1 R2 R3 53.42 53.04 53.39 AVE SD N 53.28 0.17 3.00 Table A14. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.4 (for PRMT2 + SAM, Sinefungin, or Diamine 12). A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2 vs. SAM 1.08 **** 1.4 x 10-5 PRMT2 vs. Sinefungin 1.16 **** 8.2 x 10-6 PRMT2 vs. Diamine 12 0.24 ns 0.13 Table A15. One-way ANOVA statistical testing evaluating the melting temperatures (Tm) from Figure 3.4 (for PRMT2 + SAH and/or MS023). A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2 vs. SAH 0.36 ns 0.091 PRMT2 vs. DMSO 0.50 * 0.012 PRMT2 vs. MS023 0.37 ns 0.078 PRMT2 vs. MS023 + SAH 0.51 * 0.010 SAH vs. DMSO 0.14 ns >0.99 SAH vs. MS023 0.01 ns >0.99 SAH vs. MS023 + SAH 0.15 ns >0.99 DMSO vs. MS023 0.13 ns >0.99 DMSO vs. MS023 + SAH -0.01 ns >0.99 MS023 vs. MS023 + SAH 0.14 ns >0.99 191 Table A16. Derived melting temperature for PRMT2 (Tm1 and Tm2) with ligands from Figure 4.1. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Starred values (*) were excluded due to experimental issues. [SAH] (μM) Tm1 (°C) Tm2 (°C) [H2A] (μM) Tm1 (°C) 0 R1 R2 R3 R1 R2 R3 0 R1 R2 R3 46.63 46.16 46.75 49.72 49.17 50.11 52.91 52.51 52.55 Ave SD N Ave SD N Ave SD N 46.51 0.25 3 49.67 0.39 3 52.66 0.18 3 1 R1 R2 R3 R1 R2 R3 1 R1 R2 R3 46.71 47.07* 47.16 50.43 53.03* 51.03 53.35 52.68 52.59 Ave SD N Ave SD N Ave SD N 46.93 0.23 2 50.73 0.3 2 52.87 0.34 3 5 R1 R2 R3 R1 R2 R3 5 R1 R2 R3 46.57 47.08 46.2 49.94 50.92 49.9 52.63 52.64 52.80 Ave SD N Ave SD N Ave SD N 46.62 0.36 3 50.25 0.47 3 52.69 0.08 3 10 R1 R2 R3 R1 R2 R3 10 R1 R2 R3 47 46.87 47.17 50.47 50.17 50.71 52.91 52.92 52.85 Ave SD N Ave SD N Ave SD N 47.01 0.12 3 50.45 0.22 3 52.89 0.03 3 50 R1 R2 R3 R1 R2 R3 50 R1 R2 R3 46.79 46.76 46.39 50.27 50.41 49.48 53.20 53.29 53.27 Ave SD N Ave SD N Ave SD N 46.65 0.18 3 50.05 0.41 3 53.25 0.04 3 100 R1 R2 R3 R1 R2 R3 100 R1 R2 R3 46.93 46.73 46.84 50.5 50.32 50.01 53.79 53.81 53.76 Ave SD N Ave SD N Ave SD N 46.83 0.08 3 50.28 0.2 3 53.79 0.02 3 500 R1 R2 R3 R1 R2 R3 500 R1 R2 R3 46.61 46.46 46.06 49.9 50.29 49.28 54.66 54.66 54.55 Ave SD N Ave SD N Ave SD N 46.38 0.23 3 49.82 0.41 3 54.62 0.05 3 [H3] (μM) Tm1 (°C) [H4] (μM) Tm1 (°C) 0 R1 R2 R3 0 R1 R2 R3 47.17 46.54 47.24 46.53 45.89 45.71 Ave SD N Ave SD N 46.98 0.31 3 46.04 0.35 3 1 R1 R2 R3 1 R1 R2 R3 47.1 47.62 46.41 46.49 46.55 Ave SD N Ave SD N 47.36 0.26 2 46.48 0.06 3 5 R1 R2 R3 5 R1 R2 R3 47.06 46.9 47.25 46.43 46.51 46.33 Ave SD N Ave SD N 47.07 0.14 3 46.42 0.07 3 10 R1 R2 R3 10 R1 R2 R3 47.49 47.33 47.39 46.78 46.17 46.31 Ave SD N Ave SD N 47.4 0.06 3 46.42 0.26 3 50 R1 R2 R3 50 R1 R2 R3 51.59 51.29 51.49 47.15 47.26 46.31 Ave SD N Ave SD N 51.46 0.13 3 46.9 0.42 3 100 R1 R2 R3 100 R1 R2 R3 52.03 52.27 52.13 47.09 47.57 47.46 Ave SD N Ave SD N 52.14 0.1 3 47.37 0.2 3 500 R1 R2 R3 500 R1 R2 R3 54.71 54.76 54.77 52.18 51.71 51.86 Ave SD N Ave SD N 54.75 0.03 3 51.92 0.2 3 192 Table A17. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure 4.1. A statistical level of significance cut-off <0.05 was used. SAH Tm1 (°C) Comp. ÎTm (°C) Sum. P Value H2A Tm1 (°C) Comp. ÎTm (°C) Sum. Adjusted P Value H3 Tm1 (°C) Comp. ÎTm (°C) Sum. P Value 0 vs. 1 -0.42 ns 0.7 0 vs. 1 0.22 ns 0.59 0 vs. 1 0.38 ns >0.99 0 vs. 5 -0.10 ns >0.99 0 vs. 5 0.03 ns >0.99 0 vs. 5 0.09 ns >0.99 0 vs. 10 -0.50 ns 0.23 0 vs. 10 0.24 ns 0.46 0 vs. 10 0.42 ns >0.99 0 vs. 50 -0.13 ns >0.99 0 vs. 50 0.6 ** 1.57 x 10-3 0 vs. 50 4.47 **** 8.35 x 10 -12 0 vs. 100 -0.32 ns >0.99 0 vs. 100 1.13 **** 1.58 x 10-6 0 vs. 100 5.16 **** 1.02 x 10 -12 0 vs. 500 0.140 ns >0.99 0 vs. 500 1.97 **** 1.33 x 10 -9 0 vs. 500 7.77 **** 2.00 x 10 -15 SAH Tm2 (°C) Comp. ÎTm (°C) Sum. P Value H4 Tm1 (°C) Comp. ÎTm (°C) Sum. P Value 0 vs. 1 -1.06 ns 0.13 0 vs. 1 -0.44 ns 0.74 0 vs. 5 -0.58 ns 0.87 0 vs. 5 -0.38 ns >0.99 0 vs. 10 -0.78 ns 0.32 0 vs. 10 -0.38 ns >0.99 0 vs. 50 -0.39 ns >0.99 0 vs. 50 -0.86 * 0.03 0 vs. 100 -0.61 ns 0.77 0 vs. 100 -1.33 *** 6.4 x 10 -4 0 vs. 500 -0.16 ns >0.99 0 vs. 500 -5.87 **** 8.61 x 10 -13 Table A18. Derived melting temperatures for PRMT2 (Tm1) from Figure A16. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Tm PRMT2 (ºC) PRMT2 (apo) SAH H3 SAH + H3 R1 R1 R2 R1 R1 R1 R1 R2 R3 R1 R2 R3 47.17 46.54 47.24 46.50 46.78 46.41 54.71 54.76 54.77 53.89 54.01 53.89 AVE SD N AVE SD N AVE SD N AVE SD N 46.98 0.31 3 46.56 0.15 3 54.75 0.03 3 53.93 0.06 3 193 Table A19. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A16. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2 vs. PRMT2 + SAH 0.42 ns >0.99 PRMT2 vs. PRMT2 + H3 -7.77 **** <1.00 x 10-15 PRMT2 vs. PRMT2 + SAH + H3 -6.95 **** <1.00 x 10-15 PRMT2 + SAH vs. PRMT2 + H3 -8.19 **** <1.00 x 10-15 PRMT2 + SAH vs. PRMT2 + SAH + H3 -7.37 **** <1.00 x 10-15 PRMT2 + H3 vs. PRMT2 + SAH + H3 0.82 * 0.02 Table A20. Derived melting temperature for PRMT2H112Q (Tm1) from Figure A17. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Tm (ºC) PRMT2H112Q PRMT2H112Q + SAH PRMT2H112Q + H3 PRMT2H112Q + H3 + SAH R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 43.37 43.47 43.92 44.41 43.52 43.56 43.46 42.94 44.10 43.04 42.96 44.03 Ave SD N Ave SD N Ave SD N Ave SD N 43.58 0.24 3 43.83 0.41 3 43.50 0.47 3 43.34 0.49 3 Table A21. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A17. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2H112Q vs. PRMT2H112Q + SAH -0.25 ns >0.99 PRMT2H112Q vs. PRMT2H112Q + H3 0.084 ns >0.99 PRMT2H112Q vs. PRMT2H112Q + H3 + SAH 0.24 ns >0.99 PRMT2H112Q + SAH vs. PRMT2H112Q + H3 0.33 ns >0.99 PRMT2H112Q + SAH vs. PRMT2H112Q + H3 + SAH 0.49 ns >0.99 PRMT2H112Q + H3 vs. PRMT2H112Q + H3 + SAH 0.16 ns >0.99 194 Table A22. Derived melting temperature for PRMT2ÎSH3 (Tm1) from Figure A18. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Tm (ºC) PRMT2ÎSH3 PRMT2ÎSH3 + H3 Peptide PRMT2ÎSH3 + SAH PRMT2ÎSH3 + H3 Peptide + SAH R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 40.09 39.16 39.34 41.36 42.34 42.81 40.34 40.47 39.96 41.74 42.28 43.77 AVE SD N AVE SD N AVE SD N AVE SD N 39.53 0.40 3.00 42.17 0.60 3.00 40.26 0.22 3.00 42.60 0.86 3.00 Table A23. One-way ANOVA statistical testing evaluating the for PRMT2ÎSH3 melting temperatures (Tm) from Figure A18. A statistical level of significance cut-off <0.05 was used. PRMT2ÎSH3 Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT2ÎSH3 vs. PRMT2ÎSH3 + H3 Peptide -2.64 ** 0.0029 PRMT2ÎSH3 vs. PRMT2ÎSH3 + SAH -0.73 ns 0.94 PRMT2ÎSH3 vs. PRMT2ÎSH3 + H3 Peptide + SAH -3.07 ** 0.001 PRMT2ÎSH3 + H3 Peptide vs. PRMT2ÎSH3 + SAH 1.91 * 0.021 PRMT2ÎSH3 + H3 Peptide vs. PRMT2ÎSH3 + H3 Peptide + SAH -0.43 ns >0.99 PRMT2ÎSH3 + SAH vs. PRMT2ÎSH3 + H3 Peptide + SAH -2.34 ** 0.0062 195 Table A24. Derived melting temperatures for PRMT2ÎSH3 (Tm) with H3 peptide from Figure A18. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept that corresponded with local maxima observed on the first-derivative plot. [H3 peptide] (µM) Tm PRMT2ÎSH3 (ºC) 0 R1 R2 R3 44.01 44.61 44.84 Ave SD N 44.49 0.35 3 5 R1 R2 R3 44.66 44.11 44.22 Ave SD N 44.33 0.24 3 10 R1 R2 R3 44.91 44.37 45.5 Ave SD N 44.93 0.46 3 50 R1 R2 R3 44.38 44.57 44.75 Ave SD N 44.57 0.15 3 100 R1 R2 R3 44.34 44.76 45.31 Ave SD N 44.8 0.4 3 500 R1 R2 R3 46.43 46.18 47.31 Ave SD N 46.64 0.48 3 Table A25. Derived melting temperature for PRMT1 (Tm) from Figure A19. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with local maxima observed on the first-derivative plot. Tm PRMT1 (ºC) apo H3 SAH H3 + SAH R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 49.31 48.90 49.05 50.35 50.62 50.86 52.88 53.06 53.15 57.90 57.81 57.83 AVE SD N AVE SD N AVE SD N AVE SD N 49.09 0.17 3 50.61 0.21 3 53.03 0.11 3 57.85 0.04 3 196 Table A26. Derived melting temperature for PRMT1 (Tm) with increasing amounts of H3 peptide from Figure A19. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with local maxima observed on the first-derivative plot. [H3 peptide] (μM) Tm PRMT1 (ºC) 0 R1 R2 R3 54.19 54.42 54.44 Ave SD N 54.35 0.11 3 0.5 R1 R2 R3 54.52 54.45 54.58 Ave SD N 54.51 0.05 3 1 R1 R2 R3 54.59 54.47 54.3 Ave SD N 54.45 0.12 3 5 R1 R2 R3 54.48 54.85 53.92 Ave SD N 54.42 0.38 3 10 R1 R2 R3 54.46 54.85 54.59 Ave SD N 54.63 0.16 3 50 R1 R2 R3 55.51 55.87 55.68 Ave SD N 55.69 0.14 3 100 R1 R2 R3 56.18 56.1 56.43 Ave SD N 56.24 0.14 3 500 R1 R2 R3 58.12 58.32 58.38 Ave SD N 58.28 0.11 3 197 Table A27. One-way ANOVA statistical testing evaluating the melting temperatures (Tm1) from Figure A19. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value PRMT1 vs. PRMT1 + H3 1.52 **** 4.3 x 10-6 PRMT1 vs. PRMT1 + SAH 3.94 **** 2.5 x 10-9 PRMT1 vs. PRMT1 + H3 + SAH 8.76 **** 4.2 x 10-12 Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value 0 vs. 0.5 -0.16 ns >0.99 0 vs. 1 -0.0042 ns >0.99 0 vs. 5 0.24 ns >0.99 0 vs. 10 -0.090 ns >0.99 0 vs. 50 -1.2 * 0.042 0 vs. 100 -1.8 *** 9.9 x 10-4 0 vs. 500 -3.8 **** 6.5 x 10-8 Table A28. Derived melting temperature for PRMT1/2 complex (Tm1/Tm2) from Figure 4.2. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. PRMT2 Tm (ºC) 5 μM PRMT2 Tm1 5 μM PRMT1 5 μM PRMT2 + 5 μM PRMT1 AVE SD N AVE SD N AVE SD N 45.98 0.56 3 49.34 0.17 3 49.16 0.08 3 PRMT2 + SAH Tm (ºC) 5 μM PRMT2 + 5 μM PRMT1 5 μM PRMT2 + 5 μM PRMT1 AVE SD N AVE SD N AVE SD N 46.14 0.35 3 53.28 0.12 3 53.52 0.07 3 PRMT2 + H3 peptide Tm (ºC) 5 μM PRMT2 5 μM PRMT1 5 μM PRMT2 + 5 μM PRMT1 Tm1 5 μM PRMT2 + 5 μM PRMT1 Tm2 AVE SD N AVE SD N AVE SD N AVE SD N 53.31 0.07 3 50.61 0.21 3 53.58 0.11 3 57.74 0.23 3 PRMT2 + H3 peptide + SAH Tm (ºC) 5 μM PRMT2 + H3 + SAH 5 μM PRMT1 + H3 + SAH 5 μM PRMT2 + 5 μM PRMT1 + H3 + SAH AVE SD N AVE SD N AVE SD N 55.11 0.02 3 59.83 0.04 3 58.31 0.12 3 198 Table A29. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2 from Figure 4.2. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test Tm1 Tm2 ÎTm (°C) P-value Summary apo PRMT2 vs. PRMT1/ 2 â 3.18 6.51 x 10-10 **** PRMT1 vs. PRMT1/2 â -0.18 >0.99 ns SAH PRMT2 vs. PRMT1/2 â 7.38 <1.00 x 10-15 **** PRMT1 vs. PRMT1/2 â 0.24 >0.99 ns H3 PRMT2 vs. PRMT1/2 â 3.23 2.13 x 10-11 **** PRMT1 vs. PRMT1/:2 â 0.53 0.489 ns PRMT2 vs. PRMT1/2 â -0.94 0.00364 ** PRMT1 vs. PRMT1/2 â 3.63 2.51 x 10-12 **** H3 + SAH PRMT2 vs. PRMT1/2 â 3.2 1.09 x 10-14 **** PRMT1 vs. PRMT1/2 â -1.53 1.76 x 10-6 **** 199 Table A30. Derived melting temperature for PRMT1 and PRMT2ÎSH3 from Figure 4.2. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima on the first-derivative plot. apo Tm (ºC) PRMT2ÎSH3 PRMT2ÎSH3 + PRMT1 PRMT1 R1 R2 R3 R1 R2 R3 R1 R2 R3 40.09 39.16 39.34 45.59* 47.42 47.98 49.56 49.15 49.3 AVE SD N AVE SD N AVE SD N 39.53 0.4 3 47.7 0.28 2 49.34 0.17 3 SAH Tm (ºC) PRMT2ÎSH3 PRMT2ÎSH3 + PRMT1 Tm1 PRMT1 PRMT2ÎSH3 + PRMT1 Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 40.34 40.47 39.96 53.31 53.61 53.57 53.12 53.32 53.41 63.93 64.09 63.12 AVE SD N AVE SD N AVE SD N AVE SD N 40.26 0.22 3 53.5 0.13 3 53.28 0.12 3 63.71 0.43 3 H3 peptide Tm (ºC) PRMT2ÎSH3 PRMT2ÎSH3 + PRMT1 Tm1 PRMT1 PRMT2ÎSH3 + PRMT1 Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 41.36 42.34 42.81 49.29 49.42 49.32 50.35 50.62 50.86 57.32 57.32 57.4 AVE SD N AVE SD N AVE SD N AVE SD N 42.17 0.6 3 49.34 0.06 3 50.61 0.21 3 57.35 0.04 3 H3 peptide + SAH Tm (ºC) PRMT2ÎSH3 PRMT2ÎSH3 + PRMT1 PRMT1 R1 R2 R3 R1 R2 R3 R1 R2 R3 41.74 42.28 43.77 57.37 57.36 57.37 59.89 59.8 59.81 AVE SD N AVE SD N AVE SD N 42.6 0.86 3 57.37 0.01 3 59.83 0.04 3 200 Table A31. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2ÎSH3 from Figure 4.2. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value apo PRMT2ÎSH3 vs. PRMT1/2ÎSH3 8.17 **** 5.90 x 10-8 PRMT1/2ÎSH3 vs. PRMT1 1.64 ** 2.72 x 10-4 SAH PRMT2ÎSH3 vs. PRMT1/2ÎSH3 Tm1 13.24 **** 2.32 x 10-13 PRMT2ÎSH3 vs. PRMT1/2ÎSH3 Tm2 23.45 **** <1.0 x 10-15 PRMT1/2ÎSH3 Tm1 vs. PRMT1 -0.22 ns >0.99 PRMT1/2ÎSH3 Tm2 vs. PRMT1 -10.43 **** 2.50 x 10-12 H3 peptide PRMT2ÎSH3 vs. PRMT1/2ÎSH3 Tm1 7.17 **** 7.97 x 10-10 PRMT2ÎSH3 vs. PRMT1/2ÎSH3 Tm2 15.18 **** 4.61 x 10-13 PRMT1/2ÎSH3 Tm1 vs. PRMT1 1.27 ** 5.77 x 10-3 PRMT1/2ÎSH3 Tm2 vs. PRMT1 -6.74 **** 1.47 x 10-9 H3 peptide + SAH PRMT2ÎSH3 vs. PRMT1/2ÎSH3 14.77 **** 1.65 x 10-6 PRMT1/2ÎSH3 vs. PRMT1 -2.46 ** 1.46 x 10-3 Table A32. Derived melting temperatures for PRMT1 and PRMT2H112Q from Figure 4.2. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima on the first-derivative plot. apo Tm (ºC) PRMT2H112Q PRMT1 PRMT1/2H112Q Tm1 PRMT1/2H112Q Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 43.37 43.47 43.92 49.39 49.33 49.27 42.32 42.09 41.50 50.13 50.23 50.33 AVE SD N AVE SD N AVE SD N AVE SD N 43.58 0.24 3 49.33 0.05 3 41.97 0.35 3 50.23 0.08 3 SAH Tm (ºC) PRMT2H112Q PRMT1 PRMT1/2H112Q Tm1 PRMT1/2H112Q Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 44.41 43.52 43.56 58.02 57.59 57.84 44.58 43.60 44.14 57.83 57.19 57.64 AVE SD N AVE SD N AVE SD N AVE SD N 43.83 0.41 3 57.82 0.18 3 44.11 0.40 3 57.56 0.27 3 H3 peptide Tm (ºC) PRMT2H112Q PRMT1 PRMT1/2H112Q Tm1 PRMT1/2H112Q Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 43.46 42.94 44.10 50.33 50.39 50.35 42.62 42.72 43.16 50.75 50.72 50.88 AVE SD N AVE SD N AVE SD N AVE SD N 43.50 0.47 3 50.36 0.02 3 42.83 0.24 3 50.78 0.07 3 SAH + H3 peptide Tm (ºC) PRMT2H112Q PRMT1 PRMT1/2H112Q Tm1 PRMT1/2H112Q Tm2 R1 R2 R3 R1 R2 R3 R1 R2 R3 R1 R2 R3 43.04 42.96 44.03 58.77 59.12 59.19 45.38 45.06 44.63 57.51 57.20 57.73 AVE SD N AVE SD N AVE SD N AVE SD N 43.34 0.49 3 59.02 0.18 3 45.02 0.31 3 57.48 0.22 3 201 Table A33. One-way ANOVA statistical comparison of the melting temperatures obtained for PRMT1 and PRMT2H112Q from Figure 4.2. A statistical level of significance cut-off <0.05 was used. Bonferroni's multiple comparisons test ÎTm (°C) Summary Adjusted P Value apo PRMT2H112Q vs. PRMT1/2H112Q Tm1 -1.61 ** 2.11 x 10-3 PRMT1 vs. PRMT1/2H112Q Tm2 0.90 **** 8.6 x 10-5 SAH PRMT2H112Q vs. PRMT1/2H112Q Tm1 0.28 ns >0.99 PRMT1 vs. PRMT1/2H112Q Tm2 -0.26 ns >0.99 H3 peptide PRMT2H112Q vs. PRMT1/2H112Q Tm1 -0.67 ns >0.99 PRMT1 vs. PRMT1/2H112Q Tm2 0.42 ns 0.13 H3 peptide + SAH PRMT2H112Q vs. PRMT1/2H112Q Tm1 1.68 ** 1.39 x 10-3 PRMT1 vs. PRMT1/2H112Q Tm2 -1.54 **** 5.85 x 10-8 202 Table A34. Derived melting temperature for PRMT1 and PRMT2 (at different concentrations) from Figure A23. The second derivative of fluorescence with respect to temperature was plotted and the Tm was the x-intercept which corresponded with the local maxima observed on the first-derivative plot. Tm (ºC) 0 µM PRMT1 1 µM PRMT1 5 µM PRMT1 10 µM PRMT1 0 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 48.25 48.42 48.3 49.56 49.15 49.3 50.14 50.05 50.09 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 48.32 0.07 3 49.34 0.17 3 50.1 0.03 3 1 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 43.22 43.39 42.36 46.2 46.21 45.83 48.31 48.42 48.42 52.67 52.19 53.58 49.34 49.14 49.41 49.87 50.1 50.19 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 42.99 0.45 3 46.08 0.17 3 48.38 0.05 3 52.81 0.58 3 49.3 0.11 3 50.05 0.13 4 5 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 46.64 46.03 45.27 48.6 48.73 49.1 47.32 47.44 47.01 51.59 52.89 52.94 49.23 49.05 49.2 49.81 49.95 49.73 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 45.98 0.56 3 48.81 0.21 3 47.26 0.18 3 52.47 0.63 3 49.16 0.08 3 49.83 0.09 5 10 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 48.89 48.7 49.16 52.3 51.78 51.87 47.25 46.77 46.28 54.74 54.94 55.06 48.8 48.79 49.06 52.91 53.52 53.27 49.75 50.21 50.06 54.5 54.72 54.24 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 48.92 0.19 3 51.99 0.22 3 46.77 0.39 3 54.91 0.13 3 48.88 0.12 3 53.24 0.25 3 50 0.19 6 54.49 0.19 6 203 SAH Tm (ºC) 0 µM PRMT1 1 µM PRMT1 5 µM PRMT1 10 µM PRMT1 0 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 52.25 52.47 52.53 53.12 53.32 53.41 54.22 54.16 53.92 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 52.42 0.12 3 53.28 0.12 3 54.1 0.13 3 1 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 42.7 41.64 41.48 46.02 46.36 45.72 48.18 47.92 48.21 52.53 52.16 52.58 48.73 48.92 47.99 53.31 53.59 53.22 49.84 49.83 54.27 54.25 54.57 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 41.94 0.54 3 46.03 0.26 3 48.1 0.13 3 52.42 0.19 3 48.55 0.4 3 53.37 0.16 3 49.83 0 2 54.36 0.15 3 5 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 46.61 45.78 46.02 49.66 50.07 48.94 48.45 53.04 52.64 52.57 48.35 48.16 48.32 53.43 53.61 53.54 49.61 49.41 54.23 54.19 53.82 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 46.14 0.35 3 49.86 0.2 2 48.69 0.24 2 52.75 0.21 3 48.28 0.08 3 53.52 0.07 3 49.51 0.1 2 54.08 0.18 3 10 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 49.12 48.73 48.75 51.77 51.78 51.66 54.22 54.12 54.16 53.4 53.5 53 50.3 50.91 54.57 54.43 54.33 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 48.87 0.18 3 51.74 0.06 3 54.17 0.04 3 53.3 0.22 3 50.6 0.3 2 54.45 0.1 3 204 H3 Tm (ºC) 0 µM PRMT1 1 µM PRMT1 5 µM PRMT1 10 µM PRMT1 0 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 44.52 44.64 44.41 53.86 54.15 54.37 55.39 55.46 55.31 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 44.52 0.09 3 54.12 0.21 3 55.39 0.06 3 1 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 57.2 57.13 57.13 57.07 56.92 57.15 59.49 58.93 53.91 54.13 53.97 57.58 57.63 58.04 55.32 55.36 55.12 58.94 58.98 58.37 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 57.15 0.03 3 57.05 0.09 3 59.21 0.28 2 54 0.09 3 57.75 0.21 3 55.27 0.1 3 58.76 0.28 3 5 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 56.74 56.9 56.79 54 54.02 58.87 57.5 57.61 53.47 53.73 53.55 58.02 57.74 57.45 54.46 54.74 54.64 57.91 58.86 58.11 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 56.81 0.07 3 54.01 0.01 3 57.99 0.62 2 53.58 0.11 3 57.74 0.23 3 54.61 0.12 3 58.29 0.41 3 10 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 57.05 57.27 57.11 58.89 58.45 58.56 53.31 53.06 52.84 57.51 57.79 57.7 54.11 54.36 54 58.97 58.93 58.59 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 57.14 0.09 3 58.63 0.19 3 53.07 0.19 3 57.66 0.12 3 54.16 0.15 3 58.83 0.17 3 205 H3 + SAH Tm (ºC) 0 µM PRMT1 1 µM PRMT1 5 µM PRMT1 10 µM PRMT1 0 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 55.45 54.59 55.58 59.89 59.8 59.81 64.05 63.88 63.54 59.75 59.88 59.73 67.38 67.12 66.67 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 55.21 0.44 3 59.83 0.04 3 63.82 0.21 3 59.79 0.07 3 67.06 0.29 3 1 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 55.23 55.36 55.23 55.15 55.41 55.15 59.22 59.28 59.11 67.93 68.43 67.96 59.5 59.82 59.57 68.69 68.61 68.95 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 55.27 0.06 3 55.23 0.12 3 59.2 0.07 3 68.11 0.23 3 59.63 0.14 3 68.75 0.14 3 5 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 55.1 55.1 55.13 56.27 55.98 56.03 58.33 58.44 58.15 58.87 58.65 58.65 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 55.11 0.02 3 56.09 0.12 3 58.31 0.12 3 58.72 0.1 3 10 µM PRMT2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 Tm1 Tm2 55.22 55.25 55.17 56.51 56.18 56.57 57.33 56.64 56.9 58.5 58.5 58.48 Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N Ave SD N 55.22 0.03 3 56.42 0.17 3 56.96 0.28 3 58.49 0.01 3 206 Table A35. Tabulated data from densitometry analysis in Figure A26. This data was used to plot the bar charts shown in Figures 4.4G-I. Phosphor Signal (a.u.) H3 [Histones] (mg/mL) 1:0 PRMT1:2 1:1 PRMT1:2 1:10 PRMT1:2 1:100 PRMT1:2 0.005 3430 3180 887 148 0.01 1970 1700 2210 100 0.025 1660 1830 3050 297 0.05 3340 5170 4870 811 0.075 4090 5680 4600 940 0.1 5530 5090 4560 1970 0.2 4360 4970 5900 2040 0.3 3100 4540 5060 1630 H2A [Histones] (mg/mL) 1:0 PRMT1:2 1:1 PRMT1:2 1:10 PRMT1:2 1:100 PRMT1:2 0.005 4190 4840 2360 1030 0.01 4310 4620 7010 1460 0.025 4860 3600 12200 2050 0.05 4200 4150 16200 4920 0.075 3700 6160 12200 7680 0.1 5260 2750 10200 15400 0.2 6160 4630 8260 15800 0.3 5650 1730 5190 13500 H4 [Histones] (mg/mL) 1:0 PRMT1:2 1:1 PRMT1:2 1:10 PRMT1:2 1:100 PRMT1:2 0.005 1880 2960 1800 916 0.01 2540 3860 3800 847 0.025 4560 4990 6660 70 0.05 9540 12100 10200 807 0.075 9480 11700 15000 517 0.1 15000 11200 16200 11800 0.2 16900 14200 16200 13600 0.3 16300 11500 16400 16800 207 Table A36. Tabulated data from densitometry analysis from Figure A27. This data was used for fitting purposes against linear, and nonlinear regression models to determine the kinetic parameters shown in Table 4.1. [H2A] (µM) PRMT1/2 Ratio 1:0 1:1 1:10 1:100 Phosphor Signal (a.u.) 0 1367 987 1687 4410 4510 3110 7810 6810 8010 3587 3287 2887 0.1 1787 1167 1327 3410 3810 4110 8510 9810 8110 1097 1887 2487 0.5 2587 2487 1787 5410 5010 6710 10110 10310 11510 3187 5187 5487 1 2787 2087 3987 4810 4110 6710 10210 8710 8810 5487 5287 5187 5 2487 2987 3387 9910 11410 11410 12110 15210 14110 4787 5687 4887 10 9587 9287 8887 17510 21310 19910 21610 20010 18810 8487 8687 7787 15 7787 8287 9187 33710 38610 36310 28610 33810 31910 7087 6587 7987 20 14287 15787 13487 43610 46210 39810 48510 43810 43110 10487 10487 9287 25 20187 16387 18987 52310 52210 46810 44510 44710 37110 7287 6187 6987 Table A37. Tabulated data from densitometry analysis from Figure A28. This data was used for fitting purposes against linear, and nonlinear regression models to determine the kinetic parameters shown in Table 4.1. [H3] (µM) PRMT1/2 Ratio 1:0 1:1 1:10 1:100 Phosphor Signal (a.u.) 0 1587 2127 1837 2150 1150 1940 1840 1490 2770 2467 2117 3607 0.1 2817 2697 2727 1120 1330 1750 3780 2730 3420 4207 4417 4527 0.5 3247 2957 3167 1810 1680 1740 3200 2370 2850 4267 4697 5017 1 5027 4967 5597 2160 240 760 6500 6100 2940 5297 4437 4227 5 10607 10907 11707 27300 31100 28200 12500 11500 11400 5007 4557 4487 10 14307 14607 13607 55200 58400 61400 34700 37300 31700 6907 6807 6707 15 8707 8707 8807 47900 43600 47600 32200 36600 42800 5267 5577 5937 20 10607 12607 12607 38500 37300 44000 44900 42200 40700 5887 6507 6307 25 9107 10007 9807 35100 40600 43400 51900 44400 57800 5357 6907 6067 208 Table A38. Tabulated data from densitometry analysis from Figure A29. This data was used for fitting purposes against linear, and nonlinear regression models to determine the kinetic parameters shown in Table 4.1. [H4] (µM) PRMT1/2 Ratio 1:0 1:1 1:10 1:100 Phosphor Signal (a.u.) 0 4227 7607 3607 12607 15607 11907 19007 16607 10607 40800 38100 32400 0.1 6507 8007 8507 10807 12907 11807 23007 13407 15107 31000 31700 32400 0.5 5607 7607 5487 15907 19007 13407 22107 21607 22007 45400 46900 46000 1 14907 12307 11907 8907 11607 12107 13000 13300 12500 33100 39800 29100 5 19107 19807 17607 23907 24107 20407 23300 24100 23900 36700 38400 33200 10 27007 32507 30607 33207 27807 30807 34300 26900 30700 58500 57200 59400 15 43907 39707 38507 47007 48907 34507 34900 40700 45600 48100 47900 41000 20 46807 45207 54107 46207 35007 36007 38400 44100 36600 62400 60000 53900 Table A39. Relative densitometry analysis for Figure 4.5E. Data was normalized to the signal (Coomassie stain or phosphor image intensity) from the wildtype histone H2A reaction with PRMT1 using the band corresponding to histone H2A. The column on the far right shows the normalized ratios of phosphor image signal over the corresponding Coomassie stain signal. Enzyme H2A variant % Coomassie % Phosphor Phosphor/Coomassie PRMT1 WT 100.00 100.00 1.00 R3K 43.36 23.86 0.55 R11K 61.28 84.26 1.38 R29N 109.40 109.64 1.00 PRMT1/2 WT 126.17 140.61 1.11 R3K 32.15 1.12 0.03 R11K 69.80 59.39 0.85 R29N 84.56 117.26 1.39
Thesis/Dissertation
eng
Pharmaceutical Sciences
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Frankel, Adam
Biochemical investigations of protein arginine N-methyltransferase 2
Text