Protein Arginine Methyltransferase 5 as a Driver of Lymphomagenesis
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Porsha Latrice Smith, B.S.
Biomedical Sciences Graduate Program
The Ohio State University
2016
Dissertation Committee:
Robert A. Baiocchi, M.D., Ph.D., Advisor
Gregory B. Lesinski, Ph.D.
David M. Lucas, Ph.D.
Deepa Sampath, Ph.D.
Copyrighted by
Porsha Latrice Smith
2016
Abstract
Over the past decade, it has become clear that oncogenesis is a process driven by a wide variety of triggers including gene mutations, gene amplifications, inflammation, and immune deficiency. The growing pool of data collected from whole genome and epigenome studies of both solid and blood cancers has pointed toward dysregulation of chromatin remodelers as a unique class of cancer drivers. Next generation sequencing studies of lymphomas have identified a wide array of somatic mutations affecting enzymes that regulate epigenetic control of gene expression. Lymphoma is a type of cancer that originates in secondary lymphoid organs and manifests as an outgrowth of transformed lymphocytes, or white blood cells (WBCs) in the blood. The majority of lymphoma cases can be grouped into the Non-Hodgkin’s lymphoma (NHL) subset and mainly occurs in B-cells. B-cell NHL is a heterogeneous set of cancers that would benefit from new therapies to improve patient progression-free survival. Cancers such as NHL typically present with a combination of genetic and epigenetic aberrations that contribute to the malignancy program. The epigenetic modifier protein arginine methyltransferase 5 (PRMT5) is required for B-cell transformation following Epstein-Barr virus (EBV) infection, and is overexpressed in various subsets of B-cell NHL. Based on these data we hypothesized that PRMT5 is a major driver of B-cell lymphomagenesis. To explore the role of PRMT5 in the development and progression of B-cell NHL we created
ii a small molecule inhibitors targeted to PRMT5. Using the NHL subset mantle cell lymphoma (MCL) as a model we tested the efficacy of the drug. We discovered that
PRMT5 was overexpressed in MCL primary samples and cell lines as compared to normal resting B cells. Furthermore, use of the small molecule inhibitor decreased the proliferation and viability in these cells without affecting the normal B-cells. Additionally, use of inhibitors caused G2/M cell cycle and decreased the expression of the oncogenic cell cycle proteins cyclin D1, CDK4 and c-myc but de-repressed the expression of the tumor suppressors PDCD4, C/EBPβ and ST7. Finally, Nanostring analysis confirmed the dysregulation of multiple microRNA pathways which is attenuated with PRMT5 inhibition.
To further explore the role of PRMT5 as a potential driver of lymphomagenesis, we created an Eµ-PRMT5 mouse model that overexpressed human PRMT5 in the B- lymphoid compartment of FVB/N mice. Using this model we demonstrated an increase in the incidence of lymphoma as compared to non-transgenic mice. Most of the lymphomas developed were pre-B-cell however the model also generated a number of
T-cell lymphomas. Indeed we were also able to isolate and propagate the Tg813 T-cell lymphoma cell line from a transgenic mouse tumor. Inhibition of PRMT5 in the Tg813 cell line with a small molecule inhibitor resulted in apoptosis and loss of the expression of the oncogenic proteins cyclin D1 and c-myc. Furthermore, engraftment of the 813 cell line into both SCID and FVB/N mice caused disseminated lymphoma, characterized by organomegaly and lymphoid tumors, and was lethal within 21 days post-engraftment.
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Genetic editing utilizing CRISPR/CAS9 technology and an inducible guide RNA system illustrated that genetic knockout of PRMT5 in this line led to decreased proliferation.
Taken together, these data demonstrate the role of PRMT5 as a lymphoma driver and describe a novel class of inhibitors in MCL as well as the first PRMT5 transgenic mouse model.
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This dissertation is dedicated to my grandparents Jimmie and Marjorie Smith.
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Acknowledgments
I want to first and foremost thank God for bringing me through this pivotal point in my life. I also want to thank the Biomedical Sciences Graduate Program for the opportunity to be a part of such a wonderful academic program. I want to especially thank my mentor Dr. Robert Baiocchi for giving me the opportunity to become his first official graduate student. I came to him in need of a fresh start and his unwavering guidance and support throughout my graduate career has been highly beneficial to me and I am deeply indebted to him. I also want to thank my committee members Drs. Greg Lesinski,
Dave Lucas, and Deepa Sampath for their patience, helpful suggestions and support.
Throughout my graduate career I have been fortunate to receive funding from several sources that supported my research efforts. I would like to thank the Ohio State
University for bestowing me with the Graduate Enrichment Fellowship for my first year of graduate school and, The National Institutes of Health (NIH) for bestowing me with a diversity supplement administered through the national heart, lung, and blood institute. I am especially grateful for the opportunity to have received funding from the OSU Center for Clinical and Translation Science in the form of a TL1 mentorship training fellowship for 2 years.
While here at OSU I also decided to get involved more with the student life experience. I want to thank the Council of Graduate Students (CGS) for allowing me to serve as a vi delegate and then for electing me as treasurer for the following year. My tenure with
CGS was definitely one to remember and the opportunity was invaluable to my personal development.
Several members of the Baiocchi lab past and present have been instrumental both to this body of work as well as to influencing my research career and I am so appreciative of their efforts. I want to particularly thank Dr. Fengting Yan for starting off a lot of this work and allowing me to work with her and soak up her vast expanse of knowledge. I want to also extend thanks to Dr. Lapo Alinari and John T. Patton for offering their scientific expertise to this project, Dr. Elshafa Ahmed for assisting with the mouse studies,
Hongshan Lai for designing one of the PRMT5 inhibitors and Emily Smith and Dr. Vrajesh
Karkhanis for designing the Crispr-Cas9 sgRNA. There are also many other people involved in this project that were not a part of the Baiocchi lab that I would like to acknowledge including the labs of Dr. Chenglong Li and Pui-kai Li for discovering and synthesizing our first PRMT5 inhibitor CMP5, Dr. Anjali Mishra who was always available for flow cytometry advice and Dr. Ayers who completed most of the immunohistochemistry work for the transgenic mouse model.
Of course I could not have done any of this without the prayers and encouragement from my family and friends. My family has been especially understanding when I have to miss holidays and family gatherings to in Columbus to do lab work and I’m so grateful they never held it against me. I moved to Columbus and basically knew no one so I want to especially thank the Martin family (Ricky, Vivian, Jasmyne and Richard) for taking me
vii in as one of their own. Additionally I want to thank Ms. Josie Handy, Ms. Connie Early and Ms. Jackie Parris and the rest of the Ephesus SDA church family for their support, prayers and free meals over the years. I also want to thank my West End SDA church family in Atlanta for their prayers and support as well.
I want to thank my friends for being understanding when I disappeared for weeks at a time, but still being supportive of my efforts overall and for calling and checking up on me periodically. I especially want to thank Dr. Thomas Kampfrath for checking in on me and offering words of encouragement. My friends Casey, Kendra, Lesley and Smita, for listening to my rants and doubts about grad school and continuously encouraging me to keep going and my cousin, “buddy” Kenny for staying up with me late at night while I wrote and keeping me company.
There are countless other people whom I know and don’t know that prayed for me and assisted me over the years and I couldn’t possibly list them all here but just know that I have forever grateful to you all for continually believing in me and for being on this journey with me.
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Vita
May 2002 ...... Pebblebrook High School
May 2006 ...... B.S. Chemistry, Georgia State University
2007 to present ...... Graduate Research Associate, Department
of Internal Medicine, The Ohio State
University
Publications
1. Alinari L, Mahasenan KV, Yan F, Karkhanis V, Chung J, Smith EM, Quinion C, Smith PL, Kim L, Patton JT, Lapalombella R, Yu B, Wu Y, Roy S, De Leo A, Pileri S, Agostinelli C, Ayers L, Bradner JE, Chen-Kiang S, Elemento O, Motiwala T, Majumder S, Byrd JC, Jacob S, Sif S, Li C, Baiocchi RA. “Selective inhibition of protein arginine methyltransferase 5 blocks initiation and maintenance of B-cell transformation.” Blood. 125(16), 2530–2543 (2015).
2. Yan F, Alinari L, Lustberg ME, Katherine Martin L, Cordero-Nieves HM, Banasavadi-Siddegowda Y, Virk S, Barnholtz-Sloan J, Bell EH, Wojton J, Jacob NK, Chakravarti A, Nowicki MO, Wu X, Lapalombella R, Datta J, Yu B, Gordon K, Haseley A, Patton JT, Smith PL, Ryu J, Zhang X, Mo X, Marcucci G, Nuovo G, Kwon CH, Byrd JC, Chiocca EA, Li C, Sif S, Jacob S, Lawler S, Kaur B, Baiocchi RA. “Genetic Validation of the Protein Arginine Methyltransferase PRMT5 as a Candidate Therapeutic Target in Glioblastoma.” Cancer Res. 2014 Mar 15; 74(6):1752-65. 3. Chung J, Karkhanis V, Tae S, Yan F, Smith P, Ayers LW, Agostinelli C, Pileri S, Denis GV, Baiocchi RA, Sif S. “Protein arginine methyltransferase 5 (PRMT5) inhibition induces lymphoma cell death through reactivation of the ix
retinoblastoma tumor suppressor pathway and polycomb repressor complex 2 (PRC2) silencing.” J Biol Chem. 2013 Dec 6; 288 (49):35534-47.
Fields of Study
Major Field: Biomedical Sciences Graduate Program
Area of Emphasis: Experimental Therapeutics
Graduate Interdisciplinary Specialization: Biomedical Clinical and Translational Science
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Table of Contents
Abstract ...... ii
Acknowledgments ...... vi
Vita ...... ix
Table of Contents ...... xi
List of Tables ...... xviii
List of Figures ...... xix
Chapter 1: Introduction ...... 1
Epigenetics ...... 1
DNA methylation ...... 2
MicroRNA ...... 6
Histone Modifications ...... 8
Epigenetic dysregulation in cancer ...... 26
PRMT5 ...... 27
Structure and function ...... 27
xi
PRMT5 in normal physiology ...... 29
PRMT5 and Cancer ...... 32
Regulation of PRMT5 ...... 40
Mantle Cell Lymphoma ...... 44
Lymphoma overview ...... 44
Summary of MCL ...... 45
Disease characteristics ...... 46
Staging in MCL ...... 47
Pathways altered in MCL ...... 47
Therapies for MCL ...... 49
PRMT5 as a potential driver of MCL ...... 51
Summary and hypothesis ...... 52
Chapter 2: PRMT5 plays a definitive role in the malignancy of MCL ...... 55
Introduction ...... 55
Materials and Methods ...... 56
Cell culture ...... 56
Isolating peripheral blood mononuclear cells (PBMCs) from MCL patient blood ...... 57
Isolating B cells from a leukopak ...... 58
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Protein isolation and quantification...... 58
Western Blot ...... 59
siRNA Transfection ...... 62
shRNA lentivirus preparation ...... 62
MTS Assay ...... 63
Immunofluorescence/Confocal microscopy ...... 64
Methyltransferase assay ...... 65
Annexin V/PI flow cytometry ...... 66
Real-time (RT) quantitative polymerase chain reaction (PCR) assay ...... 66
Cell cycle ...... 67
Co-immunoprecipitation (Co-IP) ...... 68
Chromatin immunoprecipitation (ChIP) ...... 69
Nanostring ...... 69
Statistical Analysis ...... 70
Results ...... 70
PRMT5 is an important oncogenic target in mantle cell lymphoma ...... 70
Development of a PRMT5 small molecule inhibitor ...... 73
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CMP5 treatment decrease cell viability and proliferation of MCL cell lines and
primary patient samples ...... 80
PRMT5 inhibition with CMP5 affects the cell cycle pathway in MCL ...... 82
PRMT5 and cyclin D1 associate in MCL ...... 87
Inhibition of PRMT5 activity does not increase the activity of the proteasome ...... 88
Inhibition of PRMT5 results in a loss of c-myc ...... 89
Inhibition of PRMT5 results in a de-repression of tumor suppressors ...... 90
Inhibition of PRMT5 perturbs microRNA networks in MCL ...... 94
Treatment with CMP5 has a synergistic effect when used in conjunction with other
anti-tumor drugs ...... 96
Discussion ...... 99
Chapter 3: PRMT5 transgenic mice develop lymphomas ...... 106
Introduction ...... 106
Materials and Methods ...... 108
Molecular cloning ...... 108
Housing of mice ...... 110
Murine B- and T-cell lymphoma transfection ...... 110
Inserting a single copy of PRMT5 into wild type FVB/N mouse ...... 110
xiv
Genotyping ...... 110
Protein isolation and quantification...... 111
Western Blot ...... 112
Real-time (RT) quantitative polymerase chain reaction (PCR) assay ...... 113
Cell Surface flow cytometry ...... 114
B, NK and T-cell sorting ...... 114
Histology ...... 115
Development of Cell Line from Transgenic Mouse Tumor ...... 115
813 cell culture ...... 116
Human PRMT5 PCR analysis ...... 116
Loxp and pBH PCR and Sanger sequencing ...... 117
Flow cytometry ...... 118
Intracellular flow cytometry ...... 119
T-cell Receptor (TCR) Analysis...... 120
Cytogenetics ...... 120
T-cell activation ...... 120
MTS Assay ...... 121
Annexin V/PI flow cytometry ...... 122
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813 engraftment ...... 122
Crispr-Cas9 PRMT5 knockout ...... 122
Statistical Analysis ...... 124
Results ...... 124
Establishment of the hPRMT5 model ...... 124
Baseline phenotypic characteristics of hPRMT5 transgenic mice ...... 130
Characteristics of disease in hPRMT5 transgenic mice ...... 132
Penetrance of model ...... 135
Molecular analysis of transgenic mice ...... 136
Isolation and creation of 813 cell line from a hPRMT5 transgenic tumor ...... 138
Immunophenotype of 813 cell line ...... 142
General characteristics of 813 cell line ...... 145
Use of a PRMT5 small molecule inhibitor in 813 cells causes changes in activation
and cytokine signaling ...... 147
813 cells cause malignant tumors in mice...... 153
Deletion of PRMT5 decrease cellular proliferation in 813 cells ...... 155
Discussion ...... 156
Chapter 4: Concluding Discussion ...... 162
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Summary ...... 162
Limitations of the study ...... 169
Future directions ...... 170
References...... 172
Appendix A: List of Abbreviations ...... 198
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List of Tables
Table 1. Summary of epigenetic modifications...... 25
Table 2. Antibodies used for western blots and immunofluorescence...... 61
Table 3. Tabular representation of PRMT5 expression in MCL stratified by MCL variant. 72
Table 4. List of microRNAs that increase at least 2 fold in 3 or more MCL cell lines when treated with CMP5 or shRNA...... 96
Table 5. List of microRNAs that decrease at least 2 fold in 3 or more MCL cell lines when treated with CMP5 or shRNA along with common genes that they target...... 97
Table 6. Antibodies used for flow cytometry...... 118
Table 7. Penetrance of hPRMT5 murine model...... 136
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List of Figures
Figure 1. PRMT5 catalyzes the symmetric dimethylation of arginine residues on histone proteins...... 27
Figure 2. Methylation by PRMT5 typically causes transcriptional silencing of genes...... 28
Figure 3. PRMT5 protein is over-expressed in mantle cell lymphoma...... 70
Figure 4. PRMT5 expression in MCL directly correlates with aggressive variants in mantle cell lymphoma...... 71
Figure 5. Knockdown of PRMT5 results in a decrease of cell proliferation...... 73
Figure 6. PRMT5 small molecule inhibitors are developed using an in silico model...... 74
Figure 7. Small molecule inhibitors of PRMT5 inhibit the proliferation of MCL cells...... 75
Figure 8. CMP3 and CMP5 selectively knockdown of symmetric-H4R3(2Me) in the MCL cell lines Jeko and Mino...... 76
Figure 9. Ability of candidate compounds to block PRMT5 methyltransferase activity. .... 77
Figure 10. CMP5 selectively inhibits the methyltransferase activity of PRMT5...... 78
Figure 11. Inhibition of PRMT5 activity with CMP5 is only toxic in malignant MCL cells. . 79
Figure 12. CMP5 treatment affects the histone mark symmetric-H4R3 (2Me) in MCL...... 80
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Figure 13. CMP5 treatment results in the activation of apoptotic pathways in the MCL cell lines Jeko and Mino...... 81
Figure 14. PRMT5 inhibition with CMP5 treatment in MCL attenuates cyclin D1 expression...... 82
Figure 15. Inhibition of PRMT5 activity in MCL results in time dependent loss of cyclin D1.
...... 83
Figure 16. Inhibition of PRMT5 with CMP5 affects the cell cycle in MCL...... 84
Figure 17. Inhibition of PRMT5 activity with CMP5 causes G2/m cell cycle arrest...... 85
Figure 18. CMP5 treatment causes changes in Rb expression in MCL cell lines...... 86
Figure 19. PRMT5 inhibition with CMP5 results in a loss of E2F target gene and protein products...... 87
Figure 20. CMP5 treatment does not affect the expression of other cyclins but decreases the expression of cdc2...... 88
Figure 21. PRMT5 and cyclin D1 associate in MCL...... 89
Figure 22. CMP5 treatment does not induce the proteasomal degradation pathway for cyclin D1...... 90
Figure 23. Proteasome inhibition does not rescue MCL cells from the effects of CMP5. . 91
Figure 24. Inhibition of PRMT5 with CMP5 results in a loss of c-myc protein expression in
Jeko and Mino cells...... 92
Figure 25. Inhibition of PRMT5 methyltransferase activity with CMP5 results in the de- repression of ST7...... 93
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Figure 26. Treatment with CMP5 in MCL cell lines increases the expression of C/EBPβ. .. 94
Figure 27. Inhibition of PRMT5 methyltransferase activity with CMP5 results in an increase in the expression of the tumor suppressor PDCD4...... 95
Figure 28. The PRMT5 inhibitor CMP5 acts synergistically with other anti-tumor drugs used to treat MCL...... 98
Figure 29. Schematic of Prmt5 vector...... 124
Figure 30. The pBH vector containing hPRMT5 can be efficiently transfected into EL-4 and WEH1 mouse T and B-cell lymphoma cell lines respectively...... 125
Figure 31. PCR analysis of the pBH vector in the original group of pups obtained after pronuclear injections of the pBH vector containing the human PRMT5 transgene reveals
5 potential founder mice...... 126
Figure 32. The 5 candidate founder mice express hPRMT5...... 127
Figure 33. The baseline immunophenotype does not differ between non-transgenic and transgenic mice...... 128
Figure 34. Prmt5 mRNA is over-expressed in the lymphocytes of transgenic mice...... 129
Figure 35. . A small subset of transgenic mice develop lymphoma-like symptoms...... 130
Figure 36. Founder 30 displays lymphocyte infiltrate in various organs and tissues as compared to an age matched non-transgenic control...... 131
Figure 37. . Immunophenotype of Founder 30...... 132
Figure 38. F1 pup (#253) from founder 24 presented with disseminated disease...... 133
Figure 39. Mouse #253 presented with an immature B-cell lymphoma...... 134
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Figure 40. hPRMT5 transgenic mice over-express PRMT5 protein...... 135
Figure 41. Tg813 mouse has lymphoma...... 137
Figure 42. Tg813 had a rapidly proliferating T-cell lymphoma...... 138
Figure 43. The 813 cell line expresses PRMT5...... 139
Figure 44. The 813 cell line expresses the original pBH vector and loxp sites...... 140
Figure 45. The 813 cell line is a mature T-cell lymphoma cell line...... 141
Figure 46. 813 cells express intracellular CD3...... 142
Figure 47. Early and late passages of the 813 cells express the Vβ17 T-cell receptor subunit...... 143
Figure 48. 813 cells express an abnormal karyotype...... 144
Figure 49. 813 cells over-express several oncogenic proteins...... 145
Figure 50. 813 cells are sensitive to PRMT5 inhibition with CMP5...... 146
Figure 51. CMP5 treatment in 813 cells results in a loss of cyclin D1 and c-myc...... 147
Figure 52. PRMT5 modestly regulates T-cell activation 813 cells...... 148
Figure 53. 813 cells produce more IL-4 than IFN-γ...... 150
Figure 54. SCID mice engrafted with 813 cells develop an aggressive T-cell lymphoma.
...... 151
Figure 55. 813 cells cause disease in SCID mice...... 152
Figure 56. 813 cells cause disease in FVB/N mice...... 153
Figure 57. 813 cells cause lymphoma in FVB/N mice...... 154
Figure 58. Immunophenotype of splenocytes from control and 813 engrafted mice...... 155
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Figure 59. FVB/N mice engrafted with 813 cells express oncogenic proteins...... 156
Figure 60. Inducible deletion of PRMT5 in 813 cells results in decreased proliferation. . 157
xxiii
Chapter 1: Introduction
Epigenetics
Epigenetics is the study of heritable changes in gene expression that occur without alterations to the underlying deoxyribonucleic acid (DNA) sequence [1]. These alterations can be post-translational modifications to the proteins in the nucleobases that comprise
DNA that occur in the form DNA methylation, changes induced by non-coding ribonucleic acid (RNA)s, variants of the histone proteins that form the structure of chromatin or post-translational modifications to the histones that comprise the histone code. The dynamic process of epigenetics is a fundamental life process that is necessary for embryogenesis and organ remodeling [2]. Despite the need for epigenetic gene regulation, the susceptibility of gene expression to environmental cues, via epigenetics, can contribute to a transformative gene expression profile. Thus, a greater understanding of the role of epigenetics in normal biology as well as in diseases like cancer will provide the opportunity to discover new drug targets for the clinic. Work over the past decade has highlighted the importance of efficient maintenance of epigenomic regulation.
Multiple next generation sequencing studies have identified mutations in chromatin remodelers and enzymes involved with epigenetic function that are associated with malignancy. Mutations involving EZH2 and DNMT enzymes have been proposed as potential drivers of leukemogenesis and lymphomagenesis, respectively [3,4]. Cancer 1 drivers are specific alterations that contribute toward the course of carcinogenesis.
Cancer drivers can include gain (EZH2) or loss (P53) of function mutations, gene amplifications (CYCLIN D1, MYC translocations), inflammation (over expression of IL6,
IL15), aberrant microRNA expression (repressed tumor suppressor miRs, over expression onco-miRs), and finally, aberrant activity of epigenetic enzymes. Here we address the dysregulation of expression of the protein arginine methyltransferase 5 (PRMT5) enzyme, first discovered to contribute toward the pathogenesis of mantle cell lymphoma. Below, the basic elements of epigenetic regulation are reviewed in the context of how epigenomic dysregulation can potentially influence the transformation of normal cells to a malignant phenotype.
DNA methylation
In most eukaryotes, the DNA molecule is comprised of two strands composed of an alternating deoxyribose sugar and phosphate backbone and four different nitrogen containing nucleobases: adenine (A), cytosine (C), guanine (G) and thymine (T) [5]. The two strands of DNA are coiled around each other to form a double helix, and the unique sequence of the four nucleobases carries the genetic information necessary for the development and normal function of an organism. In mammals, DNA methylation occurs at CpG sites or areas in the genome with a C base preceding a G base [6]. About 80% of
CpG DNA is methylated in most eukaryotes which results in strong gene silencing and is regulated by the writers that add methyl groups to the DNA, the readers that recognize
2 the methylated DNA and recruit other epigenetic modifiers and the erasers that remove methyl groups from the DNA [7].
The “writers” of methylation are the DNA methyltransferases (DNMTs) DNMT1, DNMT3A and DNMT3B. These writers recognize and methylate DNA with the consensus sequences
CG, CHG (where H stands for the base A, T, or C) and/or CHH by the addition of a methyl group at the fifth position of cytosine to form 5-methylcytosine (5mC) in eukaryotes [8].
Methylated DNA can be maintained by DNA methyltransferases and passed on to progeny though both mitosis and meiosis. DNMT1 preferentially methylates hemimethylated DNA, or DNA that is already methylated on only one strand, and is considered to be a maintenance methyltransferase that ensures that existing DNA methylation patterns are continuously propagated. It is theorized that DNMT1 ensures propagation during DNA replication by copying the methylation pattern from the parental strand onto the newly synthesized daughter strand. Although DNMT1 exclusively maintains DNA methylation patterns in normal cells, it demonstrates the ability to generate de novo DNA methylation in mammalian cancer cells [9]. In contrast, both DNMT3A and DNMT3B are considered de novo methyltransferases due to their ability to generate new DNA methylation patterns during gametogenesis, embryogenesis and potentially in somatic cells [10]. The exact mechanism of de novo methylation by
DNMT3A and DNMT3B is not clear however, it is believed to be facilitated by their ability to form collaborative dimers and to recruit and complex with chromatin remodeling enzymes, such as methyl CpG-binding domain (MBD) protein [11,12]. Additionally, the
3
DNMT3A co-factor, DNMT3-like (DNMT3L), is a catalytically inactive DNMT3 member, that increases the binding affinity of DNMT3A and DNMT3B to the methyl donor SAM and is necessary for genomic imprints in germ cells [13].
While there are relatively few “writers” of DNA methylation, the “readers” of DNA methylation consists of many members grouped into three families of proteins which includes the MBD family, zinc finger (ZnF) proteins and SET and RING finger-associated
(SRA) domain proteins. DNA methylation readers recognize and bind to methylated DNA and recruit repressive complexes that contain histone modifiers such histone deacetylases (HDACs), which remove acetyl groups from histones promoting chromatin compaction, the transcriptional repressor msin3a [14]. Finally, the “erasers” of DNA methylation are the ten-eleven translocation (TET) proteins: TET1, TET2 and TET3. The
TET proteins belong to the 2-oxoglutarate (2-OG) and Fe (II)-dioxygenase (2OGFeDO) super family, and convert 5mC to 5-hydroxymethylcytosine (5hmC), 5-formylcytosine
(5fC), or 5-carboxylcytosine (5caC). These DNA methylation intermediates are removed by either dilution through subsequent DNA replication, or by the removal of 5fC and
5caC by thymine-DNA glycosylase (TDG) with subsequent base excision repair.
Additionally, DNMT3A and DNMT3B can convert 5hmC to cytosine in an in vitro setting
[15].
As mentioned previously, DNA methylation typically occurs at specific consensus CpG sites. The exception to this are the CpG islands (CGIs), or concentrated areas of repeating
C and G bases, which are normally hypomethylated allowing for the active transcription
4 of house-keeping and tumor suppressor genes [16]. CGIs are typically located in the promoter regions of genes, which contains the transcription start site (TSS) consensus sequence necessary to start transcription of a particular gene. In the context of cancer, the opposite phenomenon is true as most CpG DNA is hypomethylated and most CGIs are hypermethylated, which results in strong silencing of tumor suppressor genes and transcriptional activation of tumor activating genes [17]. Researchers hypothesize that
DNA methylation promotes silencing by preventing the proper binding of transcription factors (TFs) due to changes in the base or through recruitment of other histone remodeling enzymes, such as HDACs by the MBD protein which causes chromatin compaction preventing transcription and silencing genes [18]. DNA methylation can also be regulated by other epigenetic processes that can either recruit or oppose DNMTs.
Specifically, histone H3 lysine 4 trimethylation (H3K4me3) prevents the binding of
DNMT3A and DNMT3L in both embryonic and somatic cells [19]. Conversely H3K9me3,
H3K27me3, and/or H3K36me3 increase the recruitment and methyltransferase activity of
DNMT3A [20]. Moreover, both the lysine methyltransferases SUV39H1 and EZH2, which are responsible for trimethylation of H3K9 and H3K27 respectively, can also directly interact with DNMT1, DNMT3A and DNMT3B. Both the SUV39H1 and EZH2 methylation marks promote DNA methylation by allowing DNMTs to localize to genes targeted for silencing [21]. Additionally H4R3me2 produced by protein arginine methyltransferase 5
(PRMT5) is a direct binding target for DNMT3A [22].
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DNA methylation can also be regulated by microRNAs. Notably, upregulation of the miR29 family in cancer cells inhibits DNMT1, DNMT3A and DNMT3B resulting in global
DNA hypomethylation in cancer cells [23,24]. Likewise, each of the DNMT family members can also be targeted by individual microRNAs. Specifically, DNMT1 can be downregulated in cholangiocarcinoma cells by mir-148a and mir-152 in response to IL-6 production [25]. While, transient expression of miR-199a-3p in teratocarcinoma cell lines targets and inhibits DNMT3A mRNA [26]. DNMT3B is downregulated by mir-145 in a feedback loop in prostate cancer cells and is also downregulated by miR-148 [27].
Additionally, long ncRNA capable of forming stem-loop structures can bind to and prevent the methylation activity DNMTs. Specifically, the extra-coding CEBPA (ecCEBPA) long ncRNA can interact with and halt the methylation activity of DNMT1 [28].
MicroRNA
Non-coding RNAs do not code for functional proteins but rather serve as post- transcriptional regulators of gene expression. Non-coding RNAs are classified into several different groups such as transcribed ultraconserved regions (T-UCRs), small nucleolar RNAs (snoRNAs), PIWI-interacting RNAs (piRNAs), large intergenic non-coding
RNAs (lincRNAs), long non-coding RNAs (lncRNAs), and microRNAs (miRs) [29]. The most well characterized group of non-coding RNAs are miRs, which are small, 20 to 22 base pair RNAs, that mediate the expression of genes post-transcriptionally by regulating the translation of mRNA into proteins. MiRs contain sequences that are complementary to their target mRNA where they bind mostly in the 3’-untranslated region (UTR) which
6 allows them modulate translation. MiRs are initially transcribed as primary microRNA
(pri-miR) with a precursor stem-loop [30]. The stem loop is cleaved in the nucleus by a complex that contains the RNase III Drosha along with its cofactor DiGeorge syndrome chromosomal [or critical] region 8 (DGCR8) to produce precursor microRNA (pre-miR).
The pre-miR is then transported to the cytoplasm by the nuclear transport receptor exportin-5 and the nuclear protein Ran-GTP where it is cleaved by Dicer into its mature miR form. The mature miR is subsequently loaded into a member of the Argonaute protein subfamily by the Dicer–TAR RNA-binding protein 2 (TARBP2 or TRBP) complex to form the RNA-induced silencing complex (RISC). The extent of silencing mediated by the
RISC complex is determined by the degree of complementarity between the miR and the target mRNA.
MicroRNAs can be located within the introns or exons of genes, or in the intergenic regions and their expression can be controlled in a variety of different ways including, amplifications, copy-number alternations, deletions, other epigenetic modifications or mutations [31]. Extronic miRs that share their promoter with the host gene are subject to the same epigenetic and other regulatory elements as the host gene, as opposed to intronic miRs that have their own promoter and as such may be regulated by different modifications than its host gene. Modifications that occur during transcription result in decreased transcription of the miR, while those that occur post-transcriptionally lead to a reduced affinity of the miR for its target mRNA. Mutations and modifications in the
Drosha and Dicer processing machinery can also affect miR expression and as expected
7 miRs are also regulated epigenetically by DNA methylation and/or histone modifications.
Similar to that which occurs for genes, hypermethylation in the promoter region silences miR expression while other histone modifications may either enhance or silence expression depending on the modification.
Histone Modifications
Chromatin forms the backbone structure of a chromosome and consists of a complex of
DNA, proteins and non-coding RNAs packaged into nucleosomes. Each nucleosome contains approximately 147 base pairs of DNA coiled around a histone core which contains two copies each of histones H2A, H2B, H3 and H4. Nucleosomes are linked to each other by a 10 – 100 base pair linker DNA region that is bound by the linker histone
H1/H5, protecting the linker DNA and its interaction with the nucleosome and prevents
ATP-dependent remodeling of chromatin [32]. Amino acid substitutions in the canonical histone protein sequence produce histone variants causing changes in the nucleosome structure which in turn alters the function of the transcriptional machinery. Histones are also susceptible to many post-translational modifications such as, ADP-ribosylation, phosphorylation, proline isomerization, sumoylation, ubiquitination, acetylation, and methylation that control gene expression by regulating the accessibility of chromatin to the transcriptional machinery [33]. Specifically, histone tail modifications stimulate chromatin relaxation into euchromatin or condensation into heterochromatin leading to transcriptional activation or repression respectively. Consequently, crosstalk between the various histone modifications results in dynamic changes in gene expression [34].
8
Histone variants
The genes that encode for canonical histone proteins do not contain introns and their mRNAs lack polyadenylated tails; instead they contain a unique 3’ stem-loop structure that stabilizes the mRNA and promotes translation [35]. Additionally, canonical histones are incorporated into chromatin in a DNA replication-dependent manner and are almost always expressed only in the S phase of the cell cycle [36]. In contrast, the genes encoding for histone variants contain introns which allows them to undergo alternative splicing and tails of their mRNAs are polyadenylated [37]. Moreover, histone variants are incorporated into chromatin, via specific histone chaperones, in a DNA replication- independent manner and can be expressed throughout all phases of the cell cycle. These differences in sequence and DNA processing result in histone variants that contain amino acid substitutions relative to the canonical histones. These amino acid differences affect their structure and in turn interactions with other histone proteins in the nucleosome. To date there have been variants identified for three of the core histones,
H2A, H2B and H3, as well as linker histone H1, with the H2A family containing the largest number of variants. H2A.Z is the most evolutionarily conserved member and has many amino acid substitutions dispersed throughout its sequence that distinguish it from canonical H2A. H2A.Z typically supports the formation of euchromatin and subsequent transcriptional activation due to destabilization of the interaction between the H2A.Z-
H2B dimer and the H3-H4 tetramer, as well as alterations in linker H1 binding [38]. As such, the H2A.Z variant tends to be clustered in nucleosomes near the TSS in gene
9 promoters and is involved in many biological processes including gene activation in developing ES cells, the recruitment of repair machinery to double-strand breaks, and overall genome stability [39]. H2A.Z is chaperoned into nucleosomes via the chromatin remodeling complexes SWR1 and INO80 [40,41]. Similar to canonical H2A, H2A.Z is also subject to post translational modifications such as ubiquitination, methylation and acetylation [42].
H2A.X is also highly conserved evolutionarily however most of its amino acid substitutions occur in the C-terminus of the sequence. Insertion of unmodified H2A.X into chromatin impairs histone H1 binding, however, phosphorylated H2A.X enhances histone H1 binding [43]. Notably, following DNA damage H2A.X is phosphorylated by ataxia telangiectasia, mutated (ATM) and ATM and Rad3-related (ATM/ATR) forming
γH2A.X which encourages chromatin relaxation to facilitate the binding and action of
DNA repair proteins [44]. After the repair is completed, γH2A.X is dephosphorylated and removed from chromatin to restart the cell cycle and restore transcription.
The variant macroH2A, named for its macro-domain containing C-terminus, is only found in mammals and represents the largest histone variant at almost three times the size of canonical H2A. There are two genes that encode for macroH2A that can undergo alternative splicing which results in 3 isoforms: macroH2A1.1, and macroH2A1.2 occur from the first gene, and macroH2A2 forms from the second gene [45,46]. MacroH2A variants differ from canonical H2A by four amino acids in the linker L1 loop. These amino acid substitutions increase the interactions between H2A-H2B dimers creating more
10 stable nucleosomes and a heterochromatin state [47]. MacroH2A is mainly found on the inactive X chromosome however it has also been found in low levels in stems cells where it is upregulated during differentiation to prevent the reprogramming of pluripotent stem cells [46,48].
H2A.B is also a mammal only variant that has only 48% sequence homology with canonical histone H2A [46]. Specifically, H2A.B lacks the last 19 C-terminal amino acids, as compared to canonical H2A, which disrupts binding within the nucleosomal octamer and prevents the binding of histone H1 [49]. As a result, H2A.B results in euchromatin, and is associated with active transcription and mRNA processing. Notably, H2A.B plays a role in spermatogenesis and is retained in mature human sperm [50].
There are two mammalian variants of H2B, both of which are associated with gametogenesis. H2B histone family member W, testis specific (H2BFWT) shares 70% sequence homology with canonical H2B, with the most variable region being a 42 amino acid extension in the N-terminal of the protein, and is only expressed in primates [51].
TH2B, shares 89% sequence homology with canonical H2B, and also has a variable N- terminal region compared to H2B that results in an increase in α-helices in the mature protein. Despite the structural differences in these variants as compared to H2B, there have been no noticeable changes in association-dissociation efficiencies with either of these variants and the H2A dimer detected in vitro or in vivo [52,53]. TH2B, so named for its specificity to the testes, replaces H2B during the spermatid elongation step in spermatogenesis and is theorized to mediate the replacement of somatic histones by
11 protamine [54]. Some TH2B expression is retained in mature sperm indicating a possible as yet undiscovered role in fertilization.
The H3 variants, H3.3 and CenH3.v, are found in all eukaryotes. In humans, H3.3 differs from canonical H3 by only 5 amino acid residues at the positions 31, 87, 89, 90, and 96
[55]. The chaperone HIRA is responsible for shuttling H3.3 to transcriptionally active sites, while death domain-associated protein (Daxx) and the chromatin remodeler α- thalassemia/mental retardation syndrome protein (ATRX) is responsible for shuttling
H3.3 to telomeres and pericentric heterochromatin [56]. Like other histones, H3.3 is subject to post-translational modifications such as methylation and acetylation and is typically associated with modifications such as H4K4me3 that result in euchromatin and transcriptional activation [57,58]. H3.3 can also mediate transcription by co-localizing with H2A.Z at the promoters of active genes [59].
CenH3, or centromere protein A (CENP-A) is a centromere-specific H3 variant that has about 50% sequence homology to canonical H3. CenH3 contains amino acid substitutions in the histone fold domain regions which allow it to target to centromeres during replication, as well as a longer N-terminal tail than H3 which further allows it to control meiosis [60]. During the G1 phase of the cell cycle, the chaperone Holliday junction recognition protein (HJURP) is responsible for shuttling CenH3 to the centromeres [61]. CenH3 is unique to all of the histone variants in that its presence in chromatin induces DNA to wrap around the nucleosomes in a right-handed orientation instead of the usual left-handed direction [62]. Phosphorylation of CenH3 is required for
12 correct mitotic progression, and both methylation and phosphorylation of CenH3 have been suggested to mediate centromeric chromatin conformation [63,64]. Additionally ubiquitination of this variant can control its deposition at heterochromatin sites [65].
In humans, linker histone H1 has 11 different subtypes/variants. H1.1 – 1.5 and H1t are considered to be subtypes rather than variants because their expression is strictly confined to the S-phase and is DNA-replication dependent much like canonical histones
[66]. Despite their low sequence homology to canonical H1 they display high sequence homology to each other and contain lysine rich C and N terminals[67]. H1.2 – 1.5 are ubiquitously expressed while H1.1 is only expressed in certain tissues and H1t is only expressed in the testis [68]. H1.0, H1.X, histone H1 testis-specific (H1T2), histone H1-like protein in speratids1 (HILS1) and histone H1 oocyte (H1oo) are true variants as they are expressed indiscriminately throughout the cell cycle and are incorporated into chromatin in a replication-independent process [66]. They also have highly variable sequences compared to H1. H1.0 accounts for up to 80% of H1 isoforms in highly differentiated cells which suggests it plays a role in the regulation of pluripotency genes during differentiation [69]. H1.X, also known as H.10, is predominantly found in less accessible chromatin domains and shuttles in and out of the nucleus during various stages of the cell cycle [70]. H1T2 and HILS1 are specific to the testis and are important in spermatogenesis. H1T2 localizes to chromatin during spermatid elongation while HILS1 helps condense chromatin during the late spermatid stage of spermatogenesis [71,72].
H1oo is important for maturation of the oocyte during meiosis and becomes
13 incorporated into sperm after fertilization where it condenses the sperm chromatin [73].
Currently it is unclear what role, if any, chaperones and/or post translational modifications play in controlling the expression and localization of H1 subtypes and variants.
Histone ADP-ribosylation
Histone ADP-ribosylation is mediated by poly (ADP-ribose) polymerases (PARPs) proteins which hydrolyze NAD+ and transfers ADP-ribose residues to histones and other proteins in response to DNA strand breaks [74]. All of the core histones as well as linker histone
H1 are affected by subjected to ADP-ribosylation. The PARP family consists of 17 members that mediate the addition of one (mono-ADP ribosylation or MAR) or more
(poly-ADP-ribosylation or PAR) ADP-ribose groups to aspartate, glutamate or lysine residues [75]. ADP-ribosylation results in the conversion of the positive charge on targeted residues into a negative charge, facilitating chromatin relaxation and transcription. ADP-ribosylation can be recognized by four different groups of proteins: macrodomains, PAR-binding zinc finger (PBZ) domain-containing proteins, PAR-binding motif (PBM) proteins and tryptophan, tryptophan, glutamate (WWE) domain-containing proteins. Macrodomain containing proteins can bind to O-acetyl-ADP-ribose (OAADPr),
MAR and PAR residues, while the PBZ domain-containing proteins: aprataxin and PNK- like factor (APLF, also called C2orf1, XIP1, or PALF), checkpoint protein with FHA and
RING domains (CHFR), and DNA cross-link repair 1A protein (DCLRE1A/SNM1A) can only bind to PAR residues [76]. Many proteins such as p53 and ATM possess the PBM motif
14 which allows them to recognize long and branched PAR chains. Conversely only E3 ubiquitin ligases and the Diphtheria toxin-like ADP-Ribosyltransferases (ARTDs) have been shown to possess the WWE domain which recognizes iso-ADP-ribose within the
PAR chain, although mutations in these proteins may allow them to recognize MAR chains [77]. As can be seen from the various members that can interact with MAR and
PAR marks, ADP-ribosylation is important for DNA repair and checkpoint regulation.
MAR and PAR, also called PARsylation, can be removed by one or more of the three classes of ADP-ribose hydrolases: PAR glycohydrolase (PARG) isoforms, ADP- ribosylhydrolases (ARHs), and the macrodomains. The best known PAR removing enzyme is PAR gylcohydrolase (PARG) which possesses endogylcohydrolase and exoglycohydrolase activity that allows it to cleave ADP-ribose chains into free chains or mono-ADP-riboses, in the case of poly-ADP-chains [78]. Alternative splicing of PARG creates various isoforms that are localized in different cellular compartments including the nucleus, cytoplasm or mitochondria. ARHs can hydrolyze the poly-ADP-ribose chain creating free ADP-ribose, however, they cannot hydrolyze the ADP-ribose-protein bond in MAR, while some macrodomain containing proteins can hydrolyze the bonds in both
PAR and MAR [76]. Since ADP-ribosylation can occur on lysine residues it competes with methylation and acetylation at these same sites. Specifically, H4K16 acetylation inhibits
ADP-ribosylation of H4. Alternatively, both mono and poly-ADP ribosylation can reduce phosphorylation of a given histone [79].
15
Histone phosphorylation
Phosphorylation involves the addition of a phosphoryl group to an amino acid group in a protein and the tails of all four histones can be phosphorylated on serine (S), threonine
(T) and tyrosine (Y) residues by protein kinases. Phosphorylation of histones is known to be most important during DNA repair; in mammalian cells, the histone variant H2A.X is phosphorylated on serine 139 (H2A.XS139ph) to create γH2A.X [44]. H2A.XS139ph is then recognized by the mediator of DNA damage checkpoint protein 1 (MDC1) which recruits repair enzymes to the repair site [80]. Once the DNA has been repaired, phosphatases such as protein phosphatase type 2A (PP2A), WPP domain interacting protein 1 (Wip1), protein phosphatase 6 (PP6) and protein phosphatase 4 (PP4), dephosphorylate γH2A.X in order to prevent the retention of repair enzymes at the damaged DNA. H2A.X can also be phosphorylated on tyrosine 142 (Y142) and this global histone modification decreases in response to DNA damage and the subsequent increase in γH2AX [81]. Phosphorylation of sites on histone H3 is typically associated with transcriptional activation however histone H3T3, S10, T11 and S28 phosphorylation have been linked to chromatin compaction indicating the necessity of cross-talk among the phosphorylation marks to regulate the dynamics of chromatin [82]. In addition to crosstalk among the phosphorylation marks on histones, histone phosphorylation also contributes to crosstalk with other histone modifications. Particularly, H3S10ph and
H3T11ph promote H3K9 and H3K14 acetylation (ac) resulting in transcriptional activation. Additionally, reading of H3S10ph mark by the 7 members of the 14-3-3
16 protein family is enhanced by H3K9ac and H3K14ac [80].Moreover, H3S28ph mediates the demethylation and acetylation of H3K27 also resulting in transcriptional activation.
Histone phosphorylation also controls transcriptional silencing and H2BK11 must be deacetylated prior to H2BS10ph mediated chromatin compaction [82]. Furthermore,
H4S1ph blocks H4ac during DNA repair or transcriptional elongation however it can cooperate with H4ac to regulate gene expression during specific cellular events.
Histone proline isomerization
The amino acid proline can take on a cis or trans conformation depending on the angle of the bond that links it to the preceding amino acid. The difference in the dihedral angle
(ω) between the cis and trans conformation is 180° which can result in substantial changes in protein folding and structure. Proline isomerization is a naturally occurring, slow, spontaneous process and the majority of proline contained in protein structures exists as the more energetically favorable trans isomer. The parvulins, cyclophilins and
FK506 binding protein (FKBPs) families are proline isomerases that can accelerate the conversion between the proline isomers up to several thousand-fold [83]. In humans the parvulin family of isomerases consists of pin1 and par14/17 which can recognize the phospho-Ser/Thr-Pro (pS/T-P) motif and catalyze the isomerization of a proline residue preceded by phosphoserine or phosphothreonine. The cyclophilin family does not have a motif preference and is made of 18 members while there are 16 FKBPs and they prefer prolines with a preceding nonpolar amino acid such as leucine [84]. Similar to other histone modification, proline isomerization can regulate and be regulated by other
17 histone modifications. Specifically, histone H3 lysine 14 acetylation (H3K14ac) increases the trans-conformation of histone H3 proline 16 (H3P16), which in turn reduces
H3K4me3 [85]. Additionally, studies with yeast have determined that the trans- conformation of H3P38 is more favorable to Set2 mediated H3K36 methylation and the addition of proline isomerases such as Pin1 increases the cis-conformation of H3P38 and prevents Set2 H3K36 methylation [86].
Histone ubiquitination
Histone proteins are the most abundantly ubiquitinated proteins in the nucleus with about 5 – 15% of H2A and 1 – 2% of H2B conjugated with ubiquitin at any given time
[87]. Ubiquitin is a 76 kDa protein that is added to lysine residues via a 3-step process
[88]. In the first step, an ubiquitin activating enzyme (E1) activates ubiquitin in an ATP- dependent reaction. In the second step, an ubiquitin-conjugating enzyme (E2) conjugates ubiquitin to a cysteine residue with a thioester bond. In the third and final step, an ubiquitin-protein isopeptide ligase (E3) transfers ubiquitin from the E2 enzyme to its targeted lysine residue [89]. Monoubiquitination of Lys 119 and Lys 123 are the most common ubiquitin markers in H2A and H2B respectively. Monoubiquitinated H2A
(H2Aub) primarily occurs in satellite regions of the genome and typically results in gene silencing while monoubiquitinated H2B (H2Bub) primarily occurs in the gene body and typically is associated with transcriptional activation of genes. Although not as common,
H3, H4 and linker H1 histones can be ubiquitinated as well. While monoubiquitination tends to serve as a signaling marker to activate or silence transcription,
18 polyubiquitination targets proteins for degradation by the 26S proteasome and is very important during the DNA damage and repair cycle. The ubiquitin ligases really interesting new gene 1B (RING1B) and H2A ubiquitin ligase/human RNA-binding RING
H2 ubiquitin-protein ligase (2A-HUB/hRUL138) mediate H2Aub at Lys 119. Likewise, the ubiquitin ligases ring finger 20 (RNF20) and RNF40 mediate H2Bub at Lys120.
Additionally, BRG1 or BRM associated factor 250b (BAF250b) can form a complex with
Elongin B and C, cullin 2 (CUL2) and RING box protein 1 (RBX1) to catalyze H2Bub, while
RNF8 and RNF168 catalyze the formation of K63-linked polyubiquitination in histones
H2A and H2AX. Ubiquitin can be recognized by various protein families with ubiquitin binding domains (UBDs) including motif interacting with ubiquitin (MIU), ubiquitin interacting motif (UIM), and UIM and MIU-related UBD (UMI) [90]. Ubiquitin is removed by proteases called deubiquitinating enzymes (DUBs). H2A specific DUBs include ubiquitin specific peptidase 16 (USP16), 2A-DUB, USP21 and BRCA associated protein 1
(BAP1) and H2B specific DUBs include ubiquitin specific protease 8 (Ubp8), Ubp10 and
USP7 [91]. DUBs act by breaking the cysteine bond between the target residue and the ubiquitin group. DUBs are essential to complete the degradation cycle as proteins targeted for degradation by polyubiquitination must be deubiquitinated prior to degradation [92]. DUBs are also necessary for the proper activation of ubiquitin- mediated transcription.
Histone ubiquitination cooperates with other histone modifications in various processes.
Particularly, H2B ubiquitination is required for the transcriptionally activating H3K4 and
19
H3K79 di- and tri-methylation marks [93]. Conversely, H2Aub can repress transcriptional activation by preventing H3K4me2 and me3, while H3K27 methylation is required for
H2Aub. Histone variants are also subject to ubiquitination and monoubiquitination of histone variant H2A.Z regulates the silencing of genes on the inactive X chromosome in female cells [94].
Histone sumoylation
Small ubiquitin-related modifier (SUMO) is an 11 kDa protein that shares 18% sequence homology with ubiquitin, although it does not have a role in degradation [95]. The
SUMO family consists of 3 members in mammals: SUMO-1, SUMO-2 (SMT3a) and
SUMO-3 (SMT3b). The SUMO group is added in an ATP-dependent manner to lysine residues in the same E1 activating-E2 conjugating-E3 ligating sequence as ubiquitin and it targets proteins with the consensus sequence γ-Lys-X-Glu, where γ is a large hydrophobic residue and X is any amino acid. Thus far the only described E1 SUMO activating enzyme is a heterodimer that consists of SUMO activating enzyme subunit 1
(SAE1) and SAE2. Additionally, the only known E2 SUMO conjugating enzyme is ubiquitin-conjugating protein 9 (UBC9). In contrast, there are several known E3-SUMO ligases including the protein inhibitor of activated stat (PIAS) family, RAN binding protein
2 (RanBP2) and the polycomb protein Pc2 [96]. Histones H2A, H2B and H3 can be weakly sumoylated however the most efficient sumoylation occurs on the five lysine residues in the tail region of histone H4 and results in transcriptional repression due to chromatin compaction [97]. Sumoylation is recognized by proteins that contain SUMO-interaction
20 motifs [90]. Sumoylation is not as common as other histone modifications and it opposes acetylation, which normally results in transcriptional activation [98]. Due to the high sequence homology sumoylation is also theorized to oppose ubiquitination however this has not yet been confirmed experimentally.
Histone acetylation
Histone acetyltransferases (HATs) mediate the transfer of acetyl groups onto lysine residues located on histone N-terminal tails. Currently there are 17 characterized HATs that can be grouped into five main families: GNAT, MYST, CBP/p300, Src and the last, as not yet named, family contains HATs that do not have any commonalities with any of the other families [99]. Acetylation decreases the positive charge on histones, reducing the interaction of the N terminus with the negatively charged phosphate groups on DNA which promotes chromatin relaxation and subsequent gene transcription [100]. The bromodomains of certain transcription factors such as bromodomain-containing protein
2 (BRD2), or BRD4, proteins containing tandem plant homeodomain (PHD) domains or other chromatin complex remodeling components such as BRG1 can interact with the acetyl groups on histone tails further promoting transcription [101]. Conversely, removal of acetyl groups by histone deacetylases (HDACs) causes transcriptional repression. The
HDAC family has 18 members plus the sirtuins and is divided into four classes (I, II, III, and IV) based on their sequence homology to the original yeast proteins [102]. The acetylation of histones H3 and H4 is strongly associated with active gene transcription.
Specifically, acetylation of H3K56, H3K64 and H3K122 at the H3-DNA interface display a
21 strong ability to disturb the electrostatic interactions in the nucleosome and mediate gene activation [103].
Histone acetylation is one of the most described histone modifications and as mentioned previously it antagonizes and/or cooperates with many of the other histone modifications to create a histone code that dynamically regulates gene expression.
Notably, histone acetylation antagonizes ADP-ribosylation, and sumolyation, while it cooperates with proline isomerization [79,85,98]. However, histone acetylation can cooperate with or antagonize phosphorylation and methylation depending on the context [82]. Of note, H3K9, H3K14 and H3K27 acetylation competes with methylation at these same locations to promote transcriptional activation in lieu of transcriptional silencing [104].
Histone methylation
Methylation of histone proteins occurs on arginine and lysine residues [105]. Protein methyltransferase (PMTs) enzymes transfer methyl groups from the methyl donor, S- adenosyl-L-methionine (SAM), to the terminal nitrogen on a ω-guanidinium side chain of target residues. Lysine methylation is regulated by protein lysine methyltransferases
(PKMTs) which mediate the addition of up to 3 methyl groups onto a targeted lysine residue. There are approximately 51 PKMTs, however only about half of these can methylate histone lysines with the remainder only able to methylate lysine residues on non-histone residues [106]. Lysine methylation can result in transcriptional activation or silencing depending on the lysine residue methylated as well as the number of methyl
22 groups added to the lysine. The key lysine methylation sites in humans are H3K4, H3K9,
H3K27, H3K36, H3K79, and H4K20 [107]. In general, H3K9me3, H3K27me3, and
H4K20me3, mediated by SUV39H1, EZH2, and SUV4-20H2 respectively, results in transcriptional silencing, while H3K4me3, H3K36me3, and H3K79, mediated by MLL1,
SETD2, and DOT1L respectively, results in transcriptional activation [108]. Lysine methylation can be recognized by a variety of proteins including PHD, chromo, tryptophan glutamate 40 (WD40), Tudor, double/tandem Tudor, malignant brain tumor
(MBT), Ankyrin repeats, zinc-finger-cysteine-tryptophan (ZF-CW) and proline, tryptophan, tryptophan, proline (PWWP) domain-containing proteins which mediate the effects of the particular lysine residue [109].
Arginine methylation is regulated by the protein arginine methyltransferases (PRMTs) which can add up to two methyl groups in an asymmetric or symmetric matter onto targeted arginine residues. PRMTs have a highly conserved core region that regulates binding to the methyl donor and substrate. All PRMTs can form intermediary monomethylation products, therefore the 9 PRMTs thus far identified PRMTs are divided into two groups based on the pattern of their dimethylation products. PRMT1, 2, 3, 4, 6
& 8 are type I PRMTs that add two methyl groups to one guanidino nitrogen atom of arginine to form ω-N(G), N(G) asymmetric dimethylation products [110]. PRMT5 and
PRMT9 are the only confirmed type II PRMTs that add one methyl group to each of the two guanidino nitrogen atoms of arginine to form ω-N(G), N(G’) symmetric dimethylation products [111]. Conflicting reports characterize PRMT7 as either a type III
23
PRMT that forms only monomethylation products or a type II PRMT that forms symmetric demethylation products, thus more studies are needed to correctly characterize this PRMT [112,113]. PRMT1 and 5 are the major type I and type II PRMTs respectively. In general, asymmetric methylation of H4R3 by PRMT1 is associated with transcriptional activation, while symmetric methylation of H4R3 by PRMT5 is associated with transcriptional repression. Similar to lysine methylation, arginine methylation is recognized by Tudor domain-containing proteins but can also be recognized by ATRX,
DNMT3, DNMT3L (ADD) domain-containing proteins [109].
There are two classes of demethylases that can remove methylation marks from histone proteins. The lysine specific histone demethylase (LSD) family is composed of two members, LSD1 and LSD2, and can only demethylate mono and di-methylated lysine residues [114]. The Jumonji C (JMJC)-domain containing family is composed of about 30 members which can be subdivided into the 6 subfamilies lysine-specific demethylase 2
(KDM2), KDM3, KDM4, KDM5, KDM6 and PHD finger family (PHF), of which only about half have been demonstrated to possess demethylase activity, however they can demethylate mono, di and tri-methylated lysine residues [115]. LSD proteins are analogs of amine oxidase that catalyze the oxidative cleavage of the α-carbon bond of a methylated lysine to form an imine intermediate, which is further hydrolyzed to form
formaldehyde, hydrogen peroxide (H2O2) and the demethylated lysine [116]. The JMJC domain demethylases are Fe (II) and 2-oxoglutarate (2-OG)-dependent dioxygenases that use a radical generated from the oxoferryl molecule to oxygenate the methyl group.
24
Epigenetic Residue Writer Reader Eraser Effect modification CG, CHG and/or MBD proteins CHH NT DNA ZnF proteins (where H DNMTs TETs Silences genes Methylation SRA domain stands for proteins the base A, T, or C) 20-22 nt Silence/induce MicroRNA Various MiRs RISC complex Various region genes Silences/induces Histone variants genes; DNA repair Macrodomains PBZ domain aspartate, ADP proteins ADP-ribosylation glutamate PARPs ribosylases, Induces genes PBM proteins lysine macrodomains and WWE domain proteins serine, 14-3-3 proteins Silences/induces Phosphorylation threonine, kinases and phosphatases genes tyrosine MDC1 parvulins, Proline Silences/induces Proline cyclophilins isomerization genes FKBPs Ubiquitin added MIU, UIM and γ-Lys-X- Ubiquitination by E1, E2 and E3 UMI domain DUBs Silences genes Glu motif ligases proteins SUMO added by Silences/induces SIM domain Sumoylation Lysine E1, E2 and E3 SUMO ligases genes proteins ligases DNA repair BRD and tandem Acetylation Lysine HATs PHD domain HDACs Induces genes proteins Tudor, PHD, Ankrykin repeats, Arginine, chromo, WD40, LSDs, Silences/induces Methylation PMTs lysine MBT, ZF-CW, JMJC domain genes PWWP and ADD domain proteins Table 1. Summary of epigenetic modifications.
The resulting unstable carbinolamine intermediate then releases formaldehyde and the demethylated lysine [117]. The only arginine demethylase described to date is the JMJC-
25 domain containing protein JMJD6 which has been shown to target and demethylate the
H3R2me2 and H4R3me2 marks catalyzed by PRMT5 [118]. The epigenetic modifications described thus far and their corresponding effects are summarized in Table 1.
Epigenetic dysregulation in cancer
As mentioned previously, the various histone modifications work together in a dynamic process to create the histone code. The histone code, along with miR and DNA methylation activity can drastically alter gene expression in a normal cell to promote malignancy as a result of environmental and genetic cues. As noted above, cancer cells demonstrate hypermethylation in CpG islands and global hypomethylation typically as a result of mutations in the DNMT enzymes themselves or the methyl group readers such as MBDs [119]. Notably changes in DNA methylation patterns can be used as prognostic events to predict outcomes in diseases such as prostate cancer and invasive breast carcinoma [120,121]. Changes in miR expression can also perpetuate cancer and a number of oncomirs have been described in both solid and hematological malignancies.
Notably miR155 is upregulated in breast cancer and oral squamous carcinoma and mediates oncogenic effects by down regulating tumor suppressors such as von Hippel-
Lindau and increasing the expression of cell cycle proteins such as cyclin D2 respectively
[122,123]. Modulations in histone modifications play critical roles in oncogenesis as well.
Particularly, histone deacetylases that remove acetyl groups from histone proteins and in turn cause transcriptional silencing of tumor suppressors are upregulated in acute myeloid leukemia (AML) [124]. Of course none of these changes are mutually exclusive
26 and often cooperate together to foster oncogenesis. Notably the protein methyltransferase protein PRMT5 associates with the SWI/SNF chromatin remodeling complex whose members often harbor gain of function mutations and play pivotal roles in cancer development [125,126]. Additionally, PRMT5 is overexpressed in mantle cell lymphoma (MCL) and knockdown of PRMT5 resulted in the de-repression of miR-92b/96
[127]. These studies implicated PRMT5 as a master epigenetic regulator and prompted our lab to explore its role in lymphomagenesis.
PRMT5
Structure and function
Figure 1. PRMT5 catalyzes the symmetric dimethylation of arginine residues on histone proteins.
PRMT5 exists as an octomer with its binding partner MEP50 which enhances its methyltransferase activity. Using the methyl donor SAM, PRMT5 symmetrically dimethylates H2AR3, H3R2, H3R8 and H4R3.
PRMT5 is a 72 kDa protein that is encoded on chromosome 14 in both humans and mice. Thus far 6 different isoforms have been described as a result of alternative splicing of Prmt5 mRNA. The catalytic domain of PRMT5 has a S-adenosyl methionine (SAM) binding domain which contains a nucleotide binding Rossmann fold and a substrate binding β-sandwich domain [128]. The active site also contains a double-E loop which contains the highly conserved glutamate residues Glu435 and Glu444. These glutamate 27
Figure 2. Methylation by PRMT5 typically causes transcriptional silencing of genes.
In the euchromatin state the chromatin is relaxed allowing DNA to be accessed by transcription factors and other epigenetic modifiers. In the heterochromatin state, the chromatin is closed preventing DNA from being accessed by transcription factors. PRMT5 typically promotes heterochromatin and subsequent gene silencing particularly of tumor suppressors
residues form a pair of salt bridges with the guanidine side chain of the substrate arginine and the ω-NG nitrogen atom methyl recipient. Like other PRMTs, PRMT5 can monomethylate and dimethylate target arginine residues using SAM as a methyl donor to add methyl groups to arginine residues. PRMT5 dimethylates arginine residues in a nonprocessive enzymatic mechanism whereby dimethylation does not proceed until the concentration of monomethylation product exceeds the concentration of the unmethylated product [129]. PRMT5 mediates gene expression by the monomethylation and symmetric dimethylation of H2AR3, H3R2, H3R8 and H4R3 [113,130,131]. The methylation activity of PRMT5 is highlighted in Figure 1 and Figure 2.
28
PRMT5 in normal physiology
PRMTs exhibit higher expression in fetal tissues relative to other mature tissue types, highlighting their roles in progenitor cell homeostasis and normal physiological development. Most of the PRMTs are localized in specific tissues however PRMT1,
PRMT4/CARM1, and PRMT5 are expressed globally in the body. In general PRMT1 and
PRMT4 activate transcription and PRMT5 silences transcription [132]. PRMT5 is localized in both the nucleus and cytoplasm in mammalian cells ensuring that it can affect a gamut of cellular processes. Global knockout of Prmt5 is embryonic lethal emphasizing its physiological importance [133]. In order to assess the role of Prmt5 in various organs, researchers have resorted to using conditional knockouts that allow for the tissue- targeted deletion of Prmt5. Selective deletion of Prmt5 in neurons leads to death of mice
14 days postnatal due to defects in neuronal cellularity and functionality [134]. Prmt5 also plays a role in gametogenesis and selective deletion of Prmt5 in primordial germ cells results in the atrophy of the seminiferous tubules of male mice, significantly smaller testes, and a lack of functional sperm as compared to wild type mice [135]. Additionally,
Prmt5 knockout results in a significant decrease in the number of ovarian germ cells due to defective meiosis [136]. The effects of loss of PRMT5 in primordial germ cells is due to the induction of the DNA damage response which causes the premature apoptosis of critical germ cells required to produce gametes [137].
In addition to histone proteins, PRMTs such as PRMT5, can methylate non-histone proteins such as RNA-binding proteins, signal transducers, DNA-binding transcriptional
29 regulators and transcriptional coregulators, which mediates their effects on normal physiological development as well as malignant growth [138]. In TNFα activated endothelial cells, PRMT5 methylates Arg140 on homeobox A9 (HOXA9) mediating TNFα induction of E selectin and other leukocyte adhesion molecules and knockout of PRMT5 with siRNA in these cells attenuates this induction. [139]. PRMT5 also methylates R30 on the p65 subunit of NFκB, which enhances the ability of nuclear factor kappa B (NFκB) to bind DNA and upregulate its target genes [140]. Additionally, PRMT5 methylation of arginine residues on the p65 subunit of NFκB regulates the expression of chemokine (C-
X-C motif) ligand 10 (CXCL10) in primary endothelial cells [141]. These studies highlight the role of PRMT5 in the chemotaxis of certain cell types a trait necessary to increase tumor invasiveness. Moreover, symmetric methylation of arginine residues on p53 by
PRMT5 stabilizes p53 expression during cellular stress allowing it to repair DNA damage and regulate p53 synthesis [142,143].
Along with adhesion and chemotaxis PRMT5 can also regulate homeostasis of nascent cells. Adult muscle stem cells (MuSC) remain in a resting, quiescent state but are able to rapidly expand to regenerate damaged muscle tissue. Prmt5 regulates the proliferation of MuSC to prevent depletion of the stem cell pool or tumor formation and loss of Prmt5 prevents the self-repair of muscle due to a lack of myogenin and loss of the ability of the
MuSC to differentiate and form myotubes [144]. In cardiac muscle PRMT5, in conjunction
with PRMT3, methylates arginine residues on the proteins that make up the NaV1.5 sodium channels increasing their density on the surface of the muscle. This results in
30 sustained action potentials at high pacing compared to control cardiomyocytes preventing cardiac hypertrophy [145].
PRMT5 also plays a role in the homeostasis of hematopoietic stem cells and mature lymphocytes. Homozygous conditional knockout of Prmt5 in two month old mice results in pancytopenia or a global decrease in white blood cells (WBCs), red blood cells (RBCs), and platelets. Specifically, loss of Prmt5 results in significant decreases in erythroid, myeloid and megakaryocyte erythroid progenitors, and any remaining progenitor cells have a defective cell cycle as a result of impaired cytokine signaling due to loss of surface cytokine receptors such as Fms-like tyrosine kinase 3 (FLT3), interleukin 6 receptor alpha chain (IL-6Ralpha) and IL-3Ralpha and a loss of quiescence [146].
Additionally in mature T-cells, PRMT5 regulates IL-2 production in response to CD3 activation and loss of PRMT5 attenuates IL-2 production [147].
Likewise, during adipogenesis, or fat cell generation, PRMT5 is recruited to the promoter of its binding partner cooperator of PRMT5 (Copr5) which facilitates its binding to histone proteins. Copr5 helps modulate adipogenic differentiation by delaying myogenic conversion and knockout of Copr5 results in deficient adipogenic conversion and loss of mRNA expression of key adipogenic differentiation regulatory genes such as Krox20, Klf4 and Klf5. Additionally, loss of Copr5 prevents the recruitment of Prmt5 and β-catenin to the Dlk-1 promoter leading to its upregulation and subsequent impairment of adipocyte differentiation [148]. PRMT5 also methylates arginine residues in the promoter of
31
PPARγ2 which is responsible for upregulating genes related to adipose-derived hormone factors [148,149].
Other studies emphasize the role of PRMT5 in regulating key cellular processes. In the golgi apparatus (GA) PRMT5 methylates arginine residues in the golgin GM130 protein mediating its expression. Loss of PRMT5 causes defects in GA ribbon formation indicating that PRMT5 plays an important function in the architectural structure of the
GA [150]. PRMT5 also works in concert with methylosome protein 50 (MEP50) to catalyze symmetric dimethylation of H3R8 and H4R3 on the promoter of the cell cycle regulator p21Cip1, decreasing p21Cip1 expression and increasing normal keratinocyte proliferation
[151]. Finally, PRMT5 modulates glial cell differentiation by suppressing the expression of the glial cell repressors Id2 and Id4 by methylating of CpG-islands in these genes [152].
These studies emphasize the importance of Prmt5 in the development and proliferation of all tissue types in the body while also highlighting its potential to cause malignancies in these tissues when dysregulated.
PRMT5 and Cancer
Solid tumors
As highlighted in the previous section, PRMT5 is required for the normal homeostasis and proliferation of many stem and nascent cell types. However, malignant cellular programs may exploit the diverse roles of PRMT5 to promote malignancy. Additionally
PRMTs such as PRMT5 can regulate the epithelial-mesenchymal transition (EMT) which promotes tumor invasiveness. Indeed, PRMT5 expression positively correlates with
32 oncogenic growth in various cancers including bladder, gastric, colorectal, lung, lymphoma and leukemia [153–155]. This section will describe the role of dysregulated
PRMT5 in many common malignancies and establish PRMT5 as a potential target in these diseases.
Glioblastoma multiforme (GBM) is the most common, and the most aggressive brain cancer in the U.S. [156]. Interestingly, in primary GBM patient-derived samples PRMT5 is overexpressed relative to normal brain tissue and a higher amount of PRMT5 directly, positively correlates with tumor grade and inversely with overall patient survival [157].
Additionally, expression of PRMT5 in GBM cell lines directly correlates with cell line growth rate and knockdown of PRMT5 with siRNA leads to cell-cycle arrest, caspase- dependent apoptosis, loss of cell migratory activity and increased expression of the tumor suppressor gene, suppressor of tumorigenicity 7 (ST7) [158]. More than 60% of
GBM cases have aberrant EGFR/ERK 1/2 signaling and knockdown of PRMT5 with shRNA, during serum starvation, lentivirus in GBM cell lines results in loss of phospho-ERK 1/2, indicating that PRMT5 plays a role in mediating ERK 1/2 signaling [157]. Furthermore, in a pre-clinical in vivo model of GBM, mice engrafted with GBM cells treated with si-
PRMT5 experience prolonged survival and less tumor burden than mice engrafted with scramble treated GBM cells [158].
Nasopharyngeal carcinoma (NPC) is the most common malignant tumor of the nasopharynx, the area behind the nose where the nasal passages and auditory tubes meet with the upper respiratory tract [159]. Although NPC is rare in the U.S. it is highly
33 prevalent in southern China. PRMT5 is overexpressed in primary NPC tissue as compared to normal adjacent tissues (NATs) and silencing PRMT5 results in a decrease in fibroblast growth factor receptor 3 (FGFR3) expression as well as increased radiosensitivity in NPC cell lines [160]. Lung neoplasias also express more PRMT5 relative to non-neoplastic lung tissues, and PRMT5 levels positively correlate with tumor grade [161]. Moreover, knockdown of PRMT5 in lung cancer cells with sh-PRMT5 prevents proliferation, causes
G1 cell cycle arrest and abolishes FGFR3, a member of the FGFR pathway that is over- activated in lung cancer [162]. Not surprisingly, aggressive lung adenocarcinomas express more cytoplasmic PRMT5 than low-grade tumors and present with an increase in
EMT, which causes epithelial cells to lose their normal contact inhibition growth program and become migratory and invasive mesenchymal stem cells [163]. Lung cancer is one of the top five most common cancers in the world and these studies implicate PRMT5 as a major transforming factor in this malignancy [156].
Breast cancer is also one of the most common cancer types in the US, with over 200,000 women and over 2,000 men newly diagnosed each year [156]. Similar to other cancers,
PRMT5 is overexpressed in breast cancer tissues relative to normal breast tissue, and increased PRMT5 expression correlates with tumor grade and worsening prognosis. In contrast to lung adenocarcinoma, PRMT5 expression is found predominantly in the nucleus of breast cancer where it is upregulated by tumor necrosis associated factor 4
(TRAF4) [164]. PRMT5 can also repress the expression of the tumor suppressor
Programmed Cell Death 4 (PDCD4) by methylating R110 likely contributing to the
34 malignancy in breast cancer [165]. Additionally, PRMT5 can methylate p53 regulating the p53 response to DNA damage in breast cancer cell lines and knockdown of PRMT5 results in a decrease in p53 stability and protein synthesis, leading to an impaired DNA damage response [143]. This same study demonstrates that PRMT5 is necessary for the expression of translation initiation factor 4e (eIF4E), a major component of the eIF4 complex that drives cap-dependent translation. Thus in addition to cell cycle regulation
PRMT5 plays an important part in the DNA damage response and protein synthesis.
Ovarian cancer is the 8th most common cancer for women in the U.S, while prostate cancer is the 2nd most common cancer for men [156,166]. Unsurprisingly, PRMT5 is highly expressed in the cytoplasm and nucleus of primary epithelial ovarian tumors as compared to normal ovaries [167]. Additionally, as expected, higher expression of PRMT5 is associated with poorer prognosis due to advanced International Federation of
Gynecology and Obstetrics (FIGO) score, poor differentiation, lymph node invasion and persistence of residual tumor. Si-RNA mediated knockdown of PRMT5 in ovarian cancer cell lines results in apoptosis due to the upregulation of the cell cycle regulator E2F1. In contrast a PRMT5-MEP50-p44 complex is highly expressed in the cytoplasm of malignant prostate tumors as opposed to benign premalignant neoplasias where it is predominantly localized in the nucleus, and forced nuclear expression of PRMT5 in prostate cancer cell lines actually inhibits proliferation [168]. Similarly in hepatocellular carcinoma (HCC), PRMT5 is significantly upregulated in malignant tissues as compared to
NATs and cytoplasmic expression, not nuclear, is negatively correlated with patient
35 survival [169]. Attenuation of PRMT5 expression in HCC cell lines with siRNA causes G-1 cell cycle arrest, prevents colony formation and causes a decrease in β-catenin protein expression. Moreover, a recent study reports that PRMT5 can methylate R31 in sterol regulatory element-binding protein 1a (SREBP1a), preventing it from being phosphorylated and targeted for degradation by the ubiquitin pathway [170]. This stabilized SREBP1a hyper-activates lipogenesis, a key transforming event activated in cancers such as HCC. HCC has a poor prognosis and represents about ¾ of liver cancer cases in the U.S thus PRMT5 could potentially be a therapeutic target for new treatments in this malignancy [156].
Colorectal cancer (CRC) is the third most common cancer in both men and women in the
United States and an estimated 93,090 and 39,610 cases of colon and rectal cancer respectively were expected to be diagnosed in the year 2015 alone [156]. PRMT5 is overexpressed in about 75% of CRC tissue as compared to NATs and nuclear PRMT5 is associated with more invasive carcinomas [156,157]. siRNA mediated knock down of
PRMT5 results in G1 arrest in CRC cell lines concomitant with a decrease in the expression of FGFR3, eIF4E, and phosphorylated AKT, ERK and mTOR, key players in the
AKT survival and growth pathway [171]. Furthermore, nude mice engrafted with CRC cells that receive intravenous si-PRMT5 fail to develop tumors as compared to mice that receive intravenous scramble, indicating that dysregulated PRMT5 is a required component in CRC establishment and progression.
36
Skin cancer is the most commonly diagnosed cancer in the U.S. however the majority of cases are benign. Despite comprising only 2% of all skin cancers, malignant melanoma makes up the majority of skin cancer deaths [156]. In malignant and metastatic melanoma, PRMT5 is upregulated and blocking its expression with siRNA results in a decrease in cell proliferation and an increase in the cell cycle regulator p27 [173].
Neuroblastoma (NB) is a cancer of the nervous system that typically occurs in children younger than 5 years of age [156]. Many aggressive neuroblastomas with poor prognoses overexpress the MYCN gene. Primary poorly differentiated NB samples overexpressing MYCN also present with strong nuclear PRMT5 staining compared to differentiated tumors without the MYCN mutation that express only cytoplasmic PRMT5
[174]. Additionally, PRMT5 and MYCN proteins associate together in representative NB cells and knockdown of PRMT5 in these cells results in apoptosis. As these studies demonstrate, PRMT5 overexpression is a driver event in many common malignancies.
Blood cancers
Since PRMT5 plays a definitive role in hematopoietic homeostasis, it is not surprising that dysregulation of PRMT5 is a hallmark in blood cancers as well. The blood cancer, leukemia is a type of cancer characterized by the uncontrolled growth of blood cells, typically WBCs, in the bone marrow. Acute myeloid leukemia (AML) is one of the most common types of leukemia and has an overall poor prognosis rate for patients regardless of age [156]. Forced over-expression of PRMT5 in AML cell lines results in an increase in cell proliferation, and colony-forming ability [154]. PRMT5 also increases the
37 mRNA and protein expression of FLT-3, a gene highly mutated in AML, and decreases the expression of mir-29b, an anti-oncomir normally repressed in AML. Additionally mice, engrafted with PRMT5 overexpressing AML cells display higher levels of circulating leukemic blasts as compared to mice engrafted with normal AML cells. Attenuation of
PRMT5 methyltransferase activity with a small molecule inhibitor reverses the proliferation and colony forming effects, decreases FLT3 expression and de-represses mir-29b.
Another blood cancer, lymphoma is characterized by an overgrowth of WBCs in the lymphatic system. In the United States, an estimated 81,080 new lymphoma cases are expected to be diagnosed in the year 2016 [156]. A greater degree of histone methylation is known to correlate with a lymphoma phenotype and in non-Hodgkin’s lymphoma PRMT5 expression, driven by various oncogenic drivers such as cyclin D1, also results in the methylation of p53 preventing p53-dependent apoptosis [175]. PRMT5 is upregulated in the non-Hodgkin’s B-cell lymphoma subsets mantle cell lymphoma
(MCL), and diffuse large B cell lymphoma (DLBCL) where it promotes the expression of
EZH2, SUZ12 and EED, members of the polycomb repressive complex 2 (PRC2). Moreover expression of the cell cycle regulator RBL2 and the pro-apoptotic proteins caspase 10
(CASP10), death associated protein (DAP1), homeo-box A5 (HOXA5), and activator of apoptosis hara-kiri (HRK) are all decreased in these cell lines. Knockdown of PRMT5 by shorthair pin RNA (sh-RNA) results in the de-repression of RBL2, CASP10, DAP1, HOXA5
38
AND HRK mRNA and protein the repression of EZH2, SUZ12 and EED and apoptotic cell death [155].
The methyltransferase ability of PRMT5 can also be exploited by other proteins to drive malignancy. Mutated cyclin D1 that is constitutively confined to the nucleus
(cyclinD1T286A) can interact with its binding partner CDK4 and direct phosphorylation of
MEP50 enhancing PRMT5 methyltransferase activity in the Eμ-D1T286A transgenic mouse model [176]. PRMT5 can then methylate the CUL4A/B ubiquitin ligases, preventing their degradation of cyclin D1. Cyclin D1 activity is then allowed to persist unchecked and to drive neoplastic growth. Additionally PRMT5-cyclinD1T286A mice develop mature T cell lymphomas [175]. Moreover, phosphorylation of MEP50-PRMT5 by mutant cyclin D1 causes p53 and E2F1 destabilizing methylation and prevents transcription of apoptotic genes such as Apaf1, Bax, Puma, Noxa, and Casp9. c-myc and
ICN driven tumors are able to promote expression of PRMT5 and upregulation of PRMT5 expression driven by Notch1 and MLL-AF9 increases p53 methylation demolishing its apoptotic properties [175].
PRMT5 mRNA and protein is overexpressed in human T-cell leukemia virus type I (HTVL-
1) positive adult T-cell leukemia/lymphoma (ATLL) cell lines, and primary peripheral blood monocytes (PBMCs) as compared to normal naïve T cells and becomes overexpressed in T-cells transformed with HTLV-1 [177]. Additionally in these cell lines,
PRMT5 and the HTLV-1 accessory protein p-30 repress viral p19 Gag production indicating that PRMT5 plays a role in balancing viral gene expression and cell survival
39 during transformation. Attenuation of PRMT5 with shRNA or a small molecule inhibitor results in the inhibition of cell proliferation and apoptosis in these cell lines as well as an increase in viral gene expression.
Regulation of PRMT5
Cofactors
For optimal methyltransferase activity PRMT5 requires the activity of its binding partner
MEP50 and human PRMT5 exists as a ~453 kDa octamer which consists of four PRMT5 proteins and four MEP50 proteins [128]. MEP50, also known as Wdr77 or androgen receptor coactivator p44, is a 7-bladed tryptophan, aspartic acid (WD) repeat β-propeller protein. The MEP50 molecules form an octamer with PRMT5 by interacting with the triosephosphate isomerase (TIM barrel) domains located in the N-terminal of PRMT5. It is theorized that the binding of MEP50 to PRMT5 increases the histone methyltransferase activity of PRMT5 by increasing its affinity for the target protein substrate [178].
In addition to and along with MEP50, PRMT5 can also associate with other protein binding partners that regulate the methyltransferase activity of PRMT5 to potentiate different effects in a variety of scenarios. As mentioned previously, the nuclear protein
COPR5 binds tightly to PRMT5 and guides it to nucleosomes where it preferentially mediates H4R3me2 residues on histones [179]. Additionally, the presence of COPR5 is required for the recruitment of PRMT5 to the promoter of genes such as the CCNE1 gene that encodes for cyclin E1 protein [180].
40
The chloride channel nucleotide sensitive 1A protein (pICln) can bind to MEP50-PRMT5, forming a pICln-MEP50-PRMT5 complex that prevents the methylation of histones while promoting symmetric demethylation of the C-terminus of the Sm proteins D1, D3 and
B/B’ [181]. sDMA of Sm D1, D3 and B/B’ increases their binding to the survival of motor neuron (SMN) complex. The SMN complex is a product of the spinal muscular atrophy gene and is necessary to ensure that the Sm proteins only assemble onto correct RNA targets and prevents their association with nonspecific RNA targets [182]. PRMT5 can also symmetrically dimethylate Arg 1810 on the polymerase (RNA) II (DNA directed) polypeptide A (POLR2A) subunit of RNA polymerase II which recruits the Tudor domain of the SMN complex [183]. SMN can then interact with the helicase senataxin and resolve the formation R-loops, or DNA-RNA hybrids that form in transcription termination sites.
This phenomenon is important and defects in this pathway may explain the neurodegeneration that occurs in diseases such as ataxia oculomoter apraxia type 2 and amyotrophic lateral sclerosis (AML).
PRMT5-MEP50 can also associate with, in a mutually exclusive pattern of binding, Rio domain containing protein (RioK1), a protein analogous to pICln [184]. RioK1 functions as an adapter protein and recruits the PRMT5-MEP50 complex to symmetrically dimethylate nucleolin. Similar to pICln and RioK1, Menin, a nuclear protein encoded by the multiple endocrine neoplasia type 1 gene (MEN1), recruits PRMT5-MEP50 to the promoters of target genes where it symmetrically dimethylates H4R3 [185,186].
41
In murine primordial germ cells the transcriptional repressor BLIMP1 recruits PRMT5 to symmetrically dimethylate H2AR3 and H4R3 to control gene regulation during embryonic development [131]. Additionally, PRMT5 controls germ cell homeostasis by sDMA of piwi proteins which can interact with small non-coding piwi-interacting RNAs
(piRNAs) [187]. The arginine methylation mark deposited by PRMT5 is recognized by
Tudor proteins which modifies piwi protein localization ultimately controlling the piRNA pathway and silencing target transposons [188]. The SNAIL corepressor protein AJUBA binds to PRMT5 to create a PRMT5-MEP50-AJUBA-SNAIL complex that can use the methyltransferase activity of PRMT5 to symmetrically dimethylate H4R3 in the promoter of the SNAIL target gene Ecadherin resulting in its transcriptional repression [189].
Most notably, PRMT5 can associate with BRG1 or hBRM, an ATPase member of the chromatin remodeling SWI/SNF complex to directly target and transcriptionally silence tumor suppressor genes such as suppressor of tumorigenicity 7 (ST7), nonmetastatic 23
(NM23), and the c-myc target gene cad by symmetric dimethylation of H3R8 and H4R3 on the promoter of these genes [130,190]. Additionally, PRMT5 association with BRG1 and the SWI/SNF complex results in the repression of the Cyp24a1 gene which encodes
for the 25-hydroxyvitamin D3 24-hydroxylase enzyme necessary for vitamin D catabolism
[191].
Demethylation
As mentioned previously, the JMJD6 protein can demethylate the PRMT5 histone marks
H4R3me2 and H3R8me2 reversing the methyltransferase activity of PRMT5 [118]. The
42 methylation mark can also be removed by peptidyl arginine deiminase (PAD) proteins which catalyze the conversion of arginine to citrulline and methylamine [192].
Specifically, the PAD4 protein can deiminate monomethyl R2, R8, R17 and R26 residues on histone H3 as well as R3 on histone H4 [192,193].
Epigenetic and post-translational modifications
PRMT5 expression itself can be controlled epigenetically, transcriptionally and post- translationally. Thus far epigenetic regulation of PRMT5 has only been described in the context of microRNA. MiR92b and MiR96 are repressed in B-cell lymphoma and re- expression of these miRs causes a decrease in PRMT5 protein [194]. Conversely, PRMT5 is transcriptionally activated by nuclear transcription factor Y (NF-Y), and knockout of NF-
Y results in a significant decrease in PRMT5 mRNA [195]. Currently identified post- translational modifications to PRMT5 include phosphorylation and ubiquitination.
Phosphorylation of PRMT5 by the tyrosine kinase JAK2V617F prevents PRMT5 from interacting with its co-factor MEP50 severely impairing its methyltransferase activity
[196]. PRMT5 can also be polyubiquitinated by the E3 ubiquitin ligase carboxyl terminus of heat shock protein 70-interacting protein (CHIP) which targets it for proteasomal degradation [197].
Small molecule inhibitors
Due to the numerous oncogenic pathways driven by PRMT5, a targeted therapy would be a welcome addition to the current arsenal of anti-oncogenic drugs either as a monotherapy or in conjunction with drugs currently on the market. To date there are a
43 few drugs that have been described that possess activity against PRMT5. The type II
PRMT inhibitor, DS-437, inhibits the symmetric dimethylation activity of both PRMT5 and
PRMT7 [198]. Recently the first of a novel class of small molecule inhibitors selectively targeted to only prevent the methylation activity of PRMT5 was described in B-cell lymphoma [199]. Treatment of normal B-cells with the PRMT5 inhibitor, CMP5, prevents
B-cell transformation by Epstein Barr Virus (EBV), and attenuates cell growth and proliferation in fully transformed B-cells while having little effect on normal healthy cells.
Epizyme, Inc has also described a selective PRMT5 inhibitor, EPZ015666, that is orally bioavailable and can prevent tumor growth in an MCL xenograft model [200].
Additionally, some bio-synthetic cyanide dyes display preferential inhibition of PRMT1 but also have limited activity against PRMT5 [201]. Overall, more work is needed to further characterize and optimize these inhibitors in various in vivo systems before clinical translation however it is foreseeable that there could be a PRMT5 inhibitor for use clinically against a variety of malignancies.
Mantle Cell Lymphoma
Lymphoma overview
An estimated 74,000 new lymphoma cases are diagnosed each year, and lymphoma is responsible for around 21,000 deaths each year. Lymphoma is divided into two main types: Hodgkin’s and non-Hodgkin’s based on morphologic and pathologic features.
Hodgkin’s lymphoma (HL) is characterized by the presence of Reed-Sternberg cells and is relatively rare, while non-Hodgkin’s lymphoma (NHL) represents approximately 90% of
44 lymphoma cases. Non-Hodgkin’s lymphoma contains many different subtypes with various prognoses and is the 6th most common cancer in the United States as of 2015
[156]. About 90% of NHL cases are of B-cell origin with the rest of the cases being of T or
NK-cell origin. NHL can be classified as centroblastic, which can be further subclassified as germinal center or non-germinal center, immunoblastic, or anaplastic morphologic variants .One type of non-Hodgkin’s lymphoma that has particularly poor prognostic rates is mantle cell lymphoma (MCL), thus these patients would benefit from an aggressive novel therapy.
Summary of MCL
Mantle cell lymphoma (MCL) represents about 6% of non-Hodgkin’s lymphomas, affects men disproportionally compared to women (3:1) and has a median age of onset of about
65. MCL results from an abnormal accumulation of naïve, mature B cells that essentially render the immune system useless. The cluster of differentiation 5 (CD5) positive B cells originate from the outer mantle zone in the lymph nodes and disseminate throughout the body taking up residence in the lymphoid tissues, bone marrow, peripheral blood and extranodal sites including those that do not normally contain lymphoid cells [202].
As a result of the abnormal accumulation of these malignant cells, affected patients present with lymphadenopathy (enlarged lymph nodes), splenomegaly (enlarged spleen), and/ or hepatomegaly (enlarged liver) [203].
Patients initially respond well to treatment, however, most usually go on to relapse.
[204]. The median survival time is 5-7 years. In highly proliferating tumors it is common
45 to find point mutations in the 3’UTR of cyclin D1 which results in shorter cyclin D1 transcripts that are more stable than the full length mRNA and lack the regulatory miR15/16 binding sites. The CDK4 locus, the binding partner of cyclin D1, is also often amplified [205]. The stabilized cyclin D1 coupled with an increase in CDK4 results in hyperactivity of the cell cycle reflecting the highly proliferative nature of the tumors.
Disease characteristics
In general MCL can be divided into 3 main variants: typical/classical, blastoid and pleomorphic [202]. Classical MCL tends to present with small to medium sized lymphocytes with irregular nuclei and unobtrusive nucleoli. Blastoid MCL has a more aggressive clinical course and tends to present with rounded nuclei, finely dispersed chromatin and unobtrusive nucleoli. Pleomorphic MCL is the most aggressive form of
MCL and presents with irregular and variably sized nuclei. The cells are B-cell in origin, and express the immunoglobulin B-cell surface markers IgM and IgD usually along with lambda of the Ig light chain. Most cases express surface cluster of CD5, normally a T-cell antigen, and CD43 expression is also highly prevalent. MCL cells also express the B-cell associated antigens such as CD20, CD22, and CD79 however they are typically CD10,
CD23 and b-cell lymphoma 6 (bcl-6) negative [202]. In addition to a cyclin D1 translocation that results in its overexpression, the over-expression of the transcription factor sex determining region-Y box 11 (SOX11) is also considered a diagnostic marker for MCL and the absence of SOX11 usually correlates with a more indolent course of disease.
46
Staging in MCL
Prior to staging a complete workup of the patient is performed which includes: a complete blood count, a bone marrow evaluation, immunophenotype of blood and bone marrow conducted by flow cytometry, a chemistry profile including lactic dehydrogenase
(LDH) levels and a computed tomography of the chest, abdomen and pelvis [206].
Evaluation of the spinal fluid may be done if the patient presents with high a Ki-67 proliferation index or the blastoid variant. After initial diagnostic procedures, the Mantle cell International Prognostic Index (MIPI) is used to group patients into risk stratification groups in order to determine the best course of therapy. Using the MIPI patients are scored based on factors indicative of shorter overall survival (OS) including: age, LDH levels, ECOG performance status and white blood count at the time of diagnosis [207].
Patients are given 0 -3 points for each prognostic factor and the points are summed to a maximum score of 11. Patients with 0 – 3 points are considered low risk with a 5-year median OS of 60%, patients with 4 -5 points are considered immediate risk with a median OS of 51 months and high risk patients with 6 – 11 points have a median OS of
29 months [208].
Pathways altered in MCL
MCL has one of the highest levels of genetic instability among lymphoid neoplasms. The hallmark feature of MCL is the t(11;14)(q13;32) translocation which results in the juxtaposition of the CCND1 gene, which encodes the cell cycle regulator cyclin D1, with the Ig heavy chain. This translocation results in the constitutive overexpression of cyclin
47
D1 which likely drives the progression of the disease. A small subset of tumors are cyclin
D1 negative and instead over-express either cyclin D2 or D3 highlighting the relevance of cell cycle dysregulation in this disease [209]. Despite the importance that cyclin D1 plays in perpetuating this disease, it is not the sole source of B-cell corruption in MCL.
Notably, transgenic mice overexpressing cyclin D1 alone do not develop MCL indicating a need for other dysregulatory genetic events to take place [210].
The cyclin-dependent kinase inhibitor 2A (CDKN2A) locus on chromosome 9 also encodes the ADP-ribosylation factor/mouse double minute 2/protein 53
(ARF/MDM2/p53) and protein 16 INK4A/cyclin-dependent kinase 4 (p16INK4A/CDK4) pathways which are also known to be disrupted in MCL [211]. Under normal conditions
INK4a inhibits CDK4 and CDK6 from phosphorylating retinoblastoma (RB) thereby maintaining an anti-proliferative state. In MCL, INK4a deletion, in conjunction with increased levels of cyclin D1 from the translocation, promotes the G1-S transition and a proliferative state. p53 is commonly mutated in MCL and ARF prevents the degradation of p53 by MDM2 thereby stabilizing the protein. Additionally, MCL cases with a wild-type
CDKN2A locus can have an overexpression of the polycomb repressive complex 1 (PRC1) member B lymphoma Mo-MLV insertion region 1 (BMI-1) [212].
Many MCL cases also demonstrate ataxia-telangiectasia (ATM) mutations demonstrating possible aberrant regulation of the DNA damage pathway. Specifically, deletions in the
11q22-23 chromosomal region result in truncated ATM proteins that are unstable and therefore are rapidly degraded. Impairment of the DNA damage pathway manifests itself
48 in a high number of chromosomal changes and polyploidy karyotypes common in most aggressive cases of MCL. Additionally, the two kinases downstream of ATM, CHK1 and
CHK2 that normally prevent cell cycle progression after ATM activation following DNA damage, may be compromised. CHK2 mutations are common and CHK2 may be downregulated in some tumors.
MCL tumors also commonly present with amplified cell survival mechanisms. Notably, the anti-apoptotic BCL2 is amplified while the pro-apoptotic BIM is deleted in some MCL cell lines. Additionally, nuclear factor kappa B (NFκB), which controls the expression of genes involved in survival and apoptotic signaling pathways, is constitutively active in primary MCL tumors and cell lines. The Akt survival pathway is also a key pathway activated in MCL [213]. Furthermore, a portion of MCL cases present with mutations in one or more epigenetic modifiers including WHSC1, MLL2, MEF2B, RB1, POT1 and
SMARCA4A.
Therapies for MCL
There is currently no agreed upon standard therapy for MCL patients and no treatment that provides a complete cure. As mentioned previously, the MIDI score is determined for each patient taking into account their age, ECOG performance status, LDH levels and white blood cell count at the time of diagnosis. Patients with a low MIDI score and that are asymptomatic are sometimes treated with a “watch and see” approach, as studies have proven there is no improvement in OS with early treatment in these patients [214].
These indolent cases represent 10 – 15% of MCL cases and may not need therapy for
49 years. For symptomatic elderly patients CHOP therapy is the standard of care and typically consists of: cyclophosphamide, an alkylating agent that damages DNA by cross- linking it, hydroxydaunorubicin, an intercalating agent, onvocin, a tubulin binding agent that prevents cells from duplicating, and prednisone, a corticosteroid. The CD20 monoclonal antibody rituximab may be added to CHOP therapy (R-CHOP) to improve the patient’s response to CHOP therapy. After a patient completes the CHOP or R-CHOP regime, rituximab maintenance (one dose every other month) is usually recommended to improve OS [215]. Elderly patients also demonstrate longer progression free survival when treated with bendamustine and rituximab both as frontline and relapse therapy.
Bendamustine has the advantage of being an alkylator and antimetabolite and it is well tolerated in these patients.
Patients who are younger or who have a lower MIDI score may benefit from and be able to tolerate more aggressive frontline therapies. Patients in this cohort display a significant progression free survival with an autologous stem cell transplant (SCT), whereby the patient’s stem cells are removed from their blood and transplanted after they undergo CHOP therapy, whole body radiation and high-dose cyclophosphamide.
CHOP therapy along with autologous SCT increases average progression free survival
(PFS) from 17 to 39 months in these patients [216]. Candidates who are not eligible for an autologous SCT may opt for other treatments R-CHOP with alternating high-dose methotrexate/cytarabine (Hyper-CVAD) [217]. Despite the high response rates, this therapy regime is highly toxic therefore a significant number of patients are unable to
50 finish the therapy. Younger patients may also receive R-HyperCVAD alternating with methotrexate plus cytosine arabinoside (MTX/Ara-C). Studies of patients receiving this therapy report an overall response rate (ORR) of 97% with a complete response rate (CR) of 87% and a median time to failure (TTF) of therapy at 4.6 years [218].
Younger patients that relapse even after receiving an autologous SCT may be considered for an allogenic SCT if a human leukocyte antigen (HLA)-matched donor can be found.
Patients may also be treated with other lines of therapy including cell cycle inhibitors such as PD0332991, proteasome inhibitors such as Bortezomib, bcl-2 family inhibitors such GX15-070, the mTOR inhibitor temsirolimus or lenalidomide. Older patients unable to tolerate any of these options may be retreated with their original frontline therapy or be considered as candidates for clinical trials testing new therapeutics [219].
PRMT5 as a potential driver of MCL
PRMT5 expression increases as B-cells become transformed by EBV, indicating that
PRMT5 is a driver of B-cell transformation and subsequent malignancy [199]. More importantly, PRMT5 is upregulated in mantle cell lymphoma (MCL) cell lines and primary patient samples relative to normal resting and activated B cells [194]. Upregulation of
PRMT5 in these cell lines results in the repression of the tumor suppressor gene suppressor of tumorigenecity 7 (ST7) due to symmetric dimethylation of H3R8 and H4R3 on the promoter of ST7. Similar to the regulation of ST7, PRMT5 can increase the symmetric dimethylation of H3R8 and H4R3 on the promoters of the RB family members
RBL1, RBL2 and RB1 in B-cell lymphoma, reducing their mRNA and protein levels,
51 preventing their inhibition of EF2 genes and promoting unchecked progression of the cell cycle [220] . Knockdown of PRMT5 with lentivirus results in de-repression of the retinoblastoma like 2 (RBL2) gene and cell cycle arrest [155]. Additionally, MCL cells display an increase in the expression of the PRC2 complex, concomitant with an increase in HDAC recruitment, that directly correlates with the upregulation of PRMT5 [155].
Moreover, attenuation of PRMT5 in MCL cell lines with shRNA results in a decrease in cyclin D1 expression and proliferation and an increase in the expression of pro-apoptotic genes and subsequent apoptosis. Additionally, EPZ015666 (GSK3235025), an orally available inhibitor of PRMT5 enzymatic activity designed by Epizyme, Inc, demonstrates dose-dependent anti-tumor activity in an in vivo xenograft MCL model. These studies implicate PRMT5 in this malignancy and the potential to use PRMT5 inhibitors as a therapy in this disease.
Summary and hypothesis
Given the heterogeneity of MCL tumors, the location of the disease in the B lymphocytes and the median age of onset, MCL remains largely incurable for most patients.
Progression free survival has improved with aggressive combination therapy however patients would still benefit from additional therapeutic agents. New therapeutics are designed to target pathways known to be dysregulated and mutated in MCL. One such potential pathway is PRMT5, which is upregulated in NHL and most malignancies. To date, inhibitors of the methyltransferase activity of PRMT5 have been effective at attenuating cell proliferation and tumor growth in in vitro and in vivo systems
52 respectively. Additionally, these drugs have been able to decrease the expression of cyclin D1, a major driver of malignancy in MCL. This dissertation will characterize the role of PRMT5 in MCL and explore its role as a driver of lymphomagenesis in a transgenic mouse model. To start, we hypothesized that PRMT5 is a driver of lymphomagenesis. To test our hypothesis we utilized the first described PRMT5 inhibitor as a tool to characterize the role of PRMT5 in in vitro and in vivo models of MCL. We were able to confirm previously published results that PRMT5 was overexpressed in MCL as compared to normal B lymphocytes, establishing the relevance of PRMT5 as a potential target in this malignancy. Moreover, inhibition of the methyltransferase activity of PRMT5 with a small molecule inhibitor attenuated proliferation and increased apoptosis in both primary MCL samples and in vitro MCL cell lines. Use of the PRMT5 inhibitor also ablated the protein expression of cyclin D1, its binding partner CDK4 and c-myc but de- repressed the expression of the tumor suppressors C/EBPβ, PDCD4 and ST7. We also developed a PRMT5 murine transgenic model that over-expressed human PRMT5 in the mouse B-lymphocytes in order to further characterize the role of PRMT5 in the development of lymphoma. Although the mouse model displayed a relatively low incidence of lymphoma, we were able to generate a T-cell lymphoma cell line from one of the mouse tumors. We were able to subsequently use this cell line for similar in vitro characterization studies that were used for the MCL cell lines, and demonstrate that
PRMT5 inhibitors could decrease the protein expression of cyclin D1 and c-myc and modulate cytokine production. We were also able to use this cell line to generate a
53 syngeneic mouse model of T-cell lymphoma. The goal of this body of work was to establish PRMT5 as a therapeutic target in lymphoma and to explore its role in B-cell transformation in this disease. The significance of this dissertation is that it extensively describes the action of a PRMT5 small molecule inhibitor in MCL and it will present data describing the first generated PRMT5 transgenic mouse model. Ultimately, these data will provide insight into the mechanism of PRMT5-driven lymphomagenesis that could potentially be applied to designing better therapeutics for clinical use.
54
Chapter 2: PRMT5 plays a definitive role in the malignancy of MCL
Introduction
Mantle cell lymphoma is an incurable form of non-Hodgkin’s lymphoma although current treatment options for MCL have extended the median overall survival from 2.5 years to around 5 years [221]. Despite the improvements in survival rates MCL is particularly challenging to treat because it is usually diagnosed in advanced stages when the lymphoma has disseminated throughout the bone marrow, lymph nodes and spleen.
Moreover, patients typically initially respond well to treatment however most eventually relapse and succumb to the disease. Due to the overall poor prognosis rate of MCL there is a need for new, more effective therapies to treat patients. The hallmark feature of MCL is the t(11; 14)(q13;q32) translocation which juxtaposes cyclin D1 onto the immunoglobulin heavy chain locus resulting in the over expression of cyclin D1. The translocation also stabilizes cyclin D1 protein by increasing its half-life cyclin D1, allowing it to remain in the nucleus, further perpetuating cell cycle disruption in this disease. The epigenetic modulator PRMT5 cooperates with oncogenic drivers such as cyclin D1 to silence genes by symmetric dimethylation of arginine residues on histone proteins
(H2AR3, H3R2, H3R8 and/or H4R3) or to modulate protein expression by symmetric dimethylation of arginine residues on the proteins themselves. In an in vivo model of
55
MCL created with a constitutively nuclear cyclin D1 mutant, D1T286A, targeted to the lymphoid compartment by the immunoglobulin enhancer Eµ, isolated tumors co-express the epigenetic modifier protein arginine methyltransferase 5 (PRMT5) along with cyclin
D1T286A complexes [176]. Additionally, lethally irradiated, syngeneic C57BL/6 mice only developed lymphoma when reconstituted with hematopoietic stem/progenitor cells
(HSPCs) transduced with PRMT5 and D1T286A retrovirus and not with HPSCs transduced with D1T286A retrovirus alone [175]. Thus PRMT5 has also been demonstrated to be over-expressed and to play an oncogenic role in many other malignancies such as glioblastoma multiforme, breast cancer, lung cancer, melanoma, nasopharyngeal carcinoma, and testicular cancer [158,160,163,164,173,222]. Based on these studies we hypothesized that over-expression of PRMT5 represents a transformative event in MCL and decided to create PRMT5-targeted inhibitors in order to explore the role of PRMT5 in MCL. We partnered with the department of Pharmacy at The Ohio State University to design and synthesize small molecule inhibitors of PRMT5. This chapter will describe the action of PRMT5 small molecule inhibitors in in vitro and in vivo models of MCL.
Materials and Methods
Cell culture
Human MCL suspension cell lines Granta, Jeko, Mino, Rec-1, SP-53, UPN-1, and Z-138 and cc-MCL (a gift from Cornell University, Ithaca, NY) were grown in RMPI without glutamine (Gibco, Grand Island, NY) supplemented with 10% heat inactivated fetal bovine serum (FBS) (Atlanta Biologicals, Norcross, GA or Sigma Aldrich, St. Louis, MO) 1X
56 glutamax (Gibco, Grand Island, NY) and 1X penicillin/streptomycin (Gibco, Grand Island,
NY). Cells were kept at 37°C in a Thermo 3110 incubator (Fisher Scientific, Pittsburgh, PA)
with 5% CO2 concentration. Cells were subcultured every 2-3 days at a concentration of 5 x 105 cells/mL. Cell counting was performed utilizing a hemocytometer and trypan blue staining to visualize dead cells.
Adherent 293T cells were grown in DMEM with glutamine (Gibco, Grand Island, NY), 10%
FBS (Gibco, Grand Island, NY), 1X penicillin/streptomycin (Gibco, Grand Island, NY). Cells were split 1:5 every 2 days when they were about 80% confluent.
Isolating peripheral blood mononuclear cells (PBMCs) from MCL patient blood
About 10 mL of blood was diluted with 20 mL of PBS pH 7.4 (Gibco, Grand Island, NY).
The 30 mL of diluted blood was layered onto 10 mL of Lymphoprep™ (STEMCELL
Technologies, Vancouver, BC, Canada) in a fresh 50 mL centrifuge tube. The layered blood was centrifuged for 30 minutes at 1500 rpm with the brake on and a slow start and stop speed. After the centrifugation the middle buffy coat layer was removed and washed with PBS pH 7.4 supplemented with 2% FBS. If necessary red blood cells were lysed using 2 mL of a red blood cell lysing buffer made with 155 mM ammonium chloride (Fisher Scientific, Pittsburgh, PA), 10 mM potassium bicarbonate (Fisher
Scientific, Pittsburgh, PA), and 0.009% EDTA pH 8.0 (Gibco, Grand Island, NY). To lyse, the cells were incubated in the lysing buffer on a shaker for 10 minutes at room temperature.
After lysing, the cells were rinsed twice with 30 mL of PBS with 2% FBS. Cells were either used immediately or frozen in freeze media composed of FBS with 10% DMSO.
57
Isolating B cells from a leukopak
Leukopaks were obtained from the American Red Cross. Blood was separated into 10 mL aliquots in 50 mL centrifuge tubes. After separating the blood, 500 µL of RosetteSep™
Human B Cell Enrichment Cocktail (STEMCELL Technologies, Vancouver, BC, Canada) was added to each 50 mL tube. The tubes were incubated for 20 minutes at room temperature with gentle agitation. After 20 minutes, the blood was diluted with 20 mL of
PBS pH 7.4 (Gibco, Grand Island, NY). The 30 mL of diluted blood was layered onto 10 mL of ficoll in a fresh 50 mL centrifuge tube. The layered blood was centrifuged for 30 minutes at 1500 rpm with the brake on and a slow start and stop speed. After the centrifugation the middle buffy coat layer was removed and washed with PBS pH 7.4 supplemented with 2% FBS. If necessary red blood cells were lysed using 2 mL of the red blood cell lysing buffer described previously. The purity of B cells was checked via flow cytometry using a CD19 (BD Biosciences San Jose, CA) antibody.
Protein isolation and quantification
Cell pellets were resuspended in RIPA buffer made with 10mM pH 7.4 Tris (Fisher
Scientific, Pittsburgh, PA) 150 mM NaCl (Fisher Scientific, Pittsburgh, PA), 1% Triton X-
100 (Fisher Scientific, Pittsburgh, PA), 1% deoxycholic acid (Acros Organics [Thermo
Fisher], Pittsburgh, PA ), 0.1% SDS (Gibco [Thermo Fisher Scientific], Grand Island, NY) and 5 mM EDTA (Fisher Scientific, Pittsburgh, PA), with the protease inhibitors Sigma and phenylmethanesufonyl fluoride (PMSF) and the phosphatase inhibitor cocktails 2 and 3
(Sigma Aldrich, St. Louis, MO) added at a 1:100 dilution. Pellets were either immediately
58 frozen at -80°C or allowed to lyse on ice for 10 minutes with vortexing before and after the incubation. After either thawing the pellets on ice or lysing on ice the samples were centrifuged at maximum speed (16,000 x g) for 10 minutes in a refrigerated 5415
Eppendorf microcentrifuge (Eppendorf, Hauppauge, NY). The cell pellets were discarded and the protein concentration of the supernatant was determined via the BCA Protein
Assay Kit (Thermo Fisher, Pittsburgh, PA) per the manufacturer’s instructions. In brief, aliquots of samples were diluted 1:10 in lysis buffer with no protease or phosphatase inhibitors and 10 µL of each were added in triplicate along with the standards to a Falcon
96-well flat bottom plate (Corning, Corning, NY). The Reagent A was mixed in a 50:1 ratio with reagent B and 200 µL of the mix was added to each well. The plate was incubated at
37°C in a cell culture incubator for 30 minutes. After 30 minutes the plate was allowed to cool on the benchtop for 10 minutes. The absorbance of the plate was read with a
Labsystems Multiskan MCC/340 (Thermo, Vantaa, Finland) absorbance plate reader.
Western Blot
20 µg of protein was combined with an equal volume of 2x laemmli sample buffer made by adding 5% β-mercaptoethanol (Fisher Scientific, Pittsburgh, PA) to a stock 2x laemmli sample buffer (Biorad, Hercules, CA). The samples were then boiled for 5 minutes at
99°C. The amount of 20 µg of protein plus 2X sample buffer was loaded into either a pre-cast TGX any kD gel (Bio-Rad, Hercules, CA) or a Tris-HCl gel made with Tris-HCL and
SDS buffer (National Diagnostics, Atlanta, GA) and 40% acrylamide (BioRad, Hercules,
CA) to the desired percentage. Precision Plus Protein ladder (Bio-Rad, Hercules, CA) was
59 loaded in the first lane of each gel to use as a reference guide for the protein weights.
The gel was allowed to run, at 200 V for a pre-cast gel or 120 V for an in-house made gel, in 1X running buffer (National Diagnostics, Atlanta, GA) until the proteins had sufficiently resolved in the gel. Separated proteins from the gel were transferred onto a
PVDF membrane pre-wet with methanol (Fisher Scientific, Pittsburgh, PA) and soaked in transfer buffer. For the precast gels the transfer buffer was made from the Transblot
Turbo transfer stock buffer (BioRad, Hercules, CA) utilizing 200 proof ethanol (Fisher
Scientific, Pittsburgh, PA). For in-house gels, transfer buffer was made with 38.6 mM glycine, 48.9 mM Tris base, 3.7% SDS and 20% methanol (Fisher Scientific, Pittsburgh,
PA). The membrane and blotting pads were soaked in transfer buffer for 15 minutes at
4°C prior to the transfer. Proteins were transferred from the gel to the membrane via the
Turboblot Plus for pre-cast gels (Bio-Rad, Hercules, CA) or TransBlot SD cell for in-house made gels (Bio-Rad, Hercules, CA). Blots were either allowed to dry for blocking or blocked with 5% milk in 1X TBST (TBS [National Diagnostics, Atlanta, GA] + 0.1% Tween
20 [Sigma Aldrich, St. Louis, MO]). Blocked blots were then incubated in primary antibody for either 2 hours at room temperature or overnight at 4°C. Primary antibody was diluted 1:1500 in 5% milk made with 1X TBST. After incubation with primary antibody, blots were rinsed four times with 1X TBST for 10 minutes each time. Blots were then incubated with secondary antibody for 1 hour at room temperature. Secondary antibody was diluted 1:4000 in 5% milk made with 1X TBST. After incubation with secondary antibody, blots were rinsed four times with 1X TBST for 10 minutes each time.
60
Protein name Company Catalog # β-actin (13E5) Cell Signaling 4970 Akt Cell Signaling 9272 pAkt S473 Cell Signaling 9271 Bax Santa Cruz sc-20067 Bcl-2 Santa Cruz sc-509
BclXL Santa Cruz sc-8392 C/EBP β (H-7) Santa Cruz sc-7962 caspase-3 Cell Signaling 9662 cdc2 (cdk1) Cell Signaling 9112 cdk2 (78B2) Cell Signaling 2546 cdk4 (DCS156) Cell Signaling 2906 cdk6 Cell Signaling 13331 c-myc Cell Signaling 9402 Cyclin A (BF683) Cell Signaling 4656 Cyclin B1 Cell Signaling 4138 Cyclin D1 Cell Signaling 2922 pCyclin D1 (Thr286) Cell Signaling 2921 Cyclin D2 (D52F9) Cell Signaling 3741 Cyclin D3 (DCS22) Cell Signaling 2936 Cyclin E (HE12) Cell Signaling 4129 E2F1 Abgent AP7593C E2F2 Santa Cruz sc-9967 ERK Cell Signaling 9102 pERK Cell Signaling 4370 GSK-3β (27C10) Cell Signaling 9315 Asym (2M3) H4R3 Active Motif 39705 sym (2Me) H4R3 Abcam 5823 HDAC2 Abcam ab12169 Mcl-1 Santa Cruz sc-819 PARP Cell Signaling 9542 p53 Santa Cruz sc-126 p65 Cell Signaling 8242 PDCD4 Cell Signaling 9535 PRMT5 Cell Signaling 2252 BD Rb Pharmingen 554144 pRb S780 Cell Signaling 8180 pRb S795 Cell Signaling 9301 pRb S803/811 Cell Signaling 8516 11945-1- ST7 Proteintech AP Table 2. Antibodies used for western blots and immunofluorescence. 61
Blots were developed utilizing either SuperSignal™ West Pico chemiluminescent substrate (Thermo Fisher, Pittsburgh, PA) or WesternBright™ ECL (Advansta, Menlo Park,
California). If a more sensitive detection method was required SuperSignal™ West Femto chemiluminescent substrate (Thermo Fisher, Pittsburgh, PA) or WesternBright™ Sirius ECL
(Advansta, Menlo Park, California) was used. Bands were visualized on Hyblot CL autoradiography film (Denville, Holliston, MA) developed by a Kodak M6B (TI-BA
Enterprises, Inc, Rochester, NY). Information for antibodies used is included in Table 2. siRNA Transfection siRNA was created from the following sequences: scramble sense AAA AAA GGG TTT TTT
TCC CCC CCT GTC TC and scramble antisense AAG GGG GTT TTC CCA AAA AAA CCT GTC
TC and PRMT5 antisense AAT GCC TAT GAA CTC TTT GCC CCT GTC TC and PRMT5 sense
AAG GCA AAG AGT TCA TAG GCA CCT GTC TC utilizing the Ambion Silencer® siRNA construction kit (Thermo Fisher Scientific, Grand Island, NY). The day prior to transfection, cells were split to 500,000 cells/mL in antibiotic free media. For the transfection, cells were electroporated with a Nucleofector® Kit and apparatus (Lonza,
Allendale, NJ) according to the kit instructions. shRNA lentivirus preparation shRNA lentivirus was generated with 293T cells. One day prior to transfection, 293T were split to 5 x 106 cells per 10 cm tissue culture plate in antibiotic-free DMEM media. The day of the transfection the media on the 293T cells was changed to fresh RPMI media.
Virus was made utilizing Opti-Mem® media and Lipofectamine® 2000 (Thermo Fisher
62
Scientific, Grand Island, NY) transfection reagent and following the procedure provided with the reagent. Plasmids for the virus were made from the following sequences, which were isolated from transformed competent bacteria and maxi-prepped (Qiagen,
Valencia, CA): Sense: GGA TCC CGT CTC AGA CAT ATG AAG TGT TTG ATA TCC GAC ACT
TCA TAT GTC TGA GAT TTT TTC CAA CTC GAG and Antisense: CTC GAG TTG GAA AAA
ATC TCA GAC ATA TGA AGT GTC GGA TAT CAA ACA CTT CAT ATG TCT GAG ACG GGA
TCC. The transfection and viral production was allowed to proceed for 72 hour before the virus was harvested for transduction. One day prior to transduction, the MCL cell lines of interest were split back to 5 x 105cells/mL in fresh RPMI media. The day of transduction, the supernatant from the 293T cells that contained the virus was filtered with 0.45 micron filter (EMD Millipore, Billerica, MA), and the cells to be transduced were concentrated to 3 x 106 cells/mL in fresh RPMI media. The cells were then plated in 6- well plates with 1 mL of cells per 1 mL of virus along with 10 µg/mL polybrene (Sigma
Aldrich, St. Louis, MO) and allowed to incubate for 72 hours after which protein and RNA were collected for subsequent analysis.
MTS Assay
Cells were split the day before to 5 x 105 cells/mL. The day of treatment 100 µL of cells were plated into a 96-well round bottom plate. 100 µL of drug diluted in media was added to the cells for a final volume of 200 µL/well. Drug concentrations were initially doubled to account for the final 1:1 dilution within the well. All treatments were performed in quadruplicate within the plate and 4 wells per plate were filled with 200 µL
63 of media only to serve as blanks. Cells were incubated for the desired time length. Cell proliferation was measured using the CellTiter 96® AQueous Non-Radioactive Cell
Proliferation Assay (Promega, Madison, WI). The MTS and PMS solution was mixed in a
20:1 ratio and 25 µL of this mixture was added to each well. The plate was allowed to incubate for 2-4 hours at 37°C. After the desired incubation time, the absorbance was read at 490 nm using a Labsystems Multiskan MCC/340 (Thermo, Vantaa, Finland) absorbance plate reader. The blank absorbance values were subtracted from the absorbance values of the samples and absorbances were graphed as a percent of the solvent (DMSO) control.
Immunofluorescence/Confocal microscopy
Glass coverslips were washed with soap, water and 70% ethanol. They were then rinsed with acetone and submerged in 0.06% poly-L-lysine (Sigma Aldrich, St. Louis, MO) for 1 hour. After the 1 hour incubation, the coverslips were rinsed twice with 2 mL of 1X PBS
(Fisher Scientific, Pittsburgh, PA). Cells of interest were split to 1 x 104 cells in 2mL and allowed to adhere to the coverslips in 6 well plates overnight. The next day, the cells attached to the coverslips were washed with 1X PBS and fixed with 4% paraformaldehyde for 15 minutes at room temperature. After the incubation, the cells were washed three times with 1X PBS and treated with 0.1% triton X-100 (Fisher
Scientific, Pittsburgh, PA) for 10 minutes at room temperature. Cells were then blocked in
10% goat serum (Sigma Aldrich, St. Louis, MO) diluted in 1X PBS for 2 hours. Meanwhile the 500 µL of the antibodies of interest were incubated with 50 µL of GST-agarose beads
64 on a rotator for 2 hours at 4°C to remove non-specific binding proteins. After the 2 hour incubation, the antibodies were centrifuged to pellet the beads and the supernatant was used for the next step. The cells were washed twice with 1X PBS and incubated with 40
µL of either preimmune serum or the primary antibody cleared previously with beads at
37°C for 2 hours. Cells were then washed three times with 1X PBS then incubated with 40
µL of FITC-labeled goat anti-rabbit secondary antibody (Sigma Aldrich, St. Louis, MO) for
1 hour at 37°C. Cells were washed for three times with 1X PBS then the nuclei were stained with Draq 5 (Cell Signaling Technology, Danvers, MA) for 5 minutes at room temperature. Afterwards the coverslips were mounted with prolong antifade reagent
(Molecular Probes, Inc., [Thermo Fisher Scientific] Pittsburgh, PA). Proteins were visualized with a fluorescence microscope using a Zeiss axioscope (Zeiss, Thornwood,
NY) at 100X magnification.
Methyltransferase assay
Compounds showing selective loss of the PRMT5 epigenetic mark were evaluated in histone methyltransferase activity assays using affinity purified recombinant PRMT1 (500 ng), PRMT4 (500 ng) (EMD Millipore, Billerica, MA), hSWI/SNF-associated PRMT5 (15 l), and hSWI/SNF-associated PRMT7 (15 l) which were incubated with 2 g of HeLa S3 core histones H3 or H4 (Roche, Indianapolis, IN) in the presence and absence of DMSO, and candidate PRMT5 inhibitors. 2 mCi of S-adenosyl-L-methyl-3H-methionine [3H]SAM
(Amersham Biosciences, Amersham, UK). Histone methyltransferase buffer was made with 25 mM NaCl, 25 mM Tris, pH 8.8. Reaction mixtures were spotted on Whatman P-81
65 filter paper, washed five times with 10 ml of 0.1 mM sodium carbonate buffer (pH 9.0) to remove unincorporated [3H]SAM, and methylated peptides were detected by scintillation counting.
Annexin V/PI flow cytometry
An equal amount of cells for each condition were pelleted by centrifugation for 5 minutes at 1500 rpm. After centrifugation the supernatant was removed and the cells were resuspended in 100 µL of a master mix containing 100 µL 1X Annexin Binding
Buffer (BD Biosciences, San Jose, CA) with 5 µL FITC Annexin V (BD Biosciences, San Jose,
CA) and 5 µL propidium iodide (BD Biosciences, San Jose, CA). The cells were allowed to stain for 15 minutes. After 15 minutes, an additional 400 µL of 1X Annexin Binding Buffer was added to the cells. Cell viability was then checked via a FC500 Beckman Coulter flow cytometer (Beckman Coulter, Pasadena, CA).
Real-time (RT) quantitative polymerase chain reaction (PCR) assay
Total RNA was prepared from cells using TRIzol reagent (Thermo Fisher Scientific, Grand
Island, NY) according to the manufacturer's instructions. cDNA was synthesized using the
Taqman MircoRNA Reverse Transcriptase Kit (Thermo Fisher Scientific, Grand Island, NY) per the kit’s directions. Real-time PCR was performed in triplicate for each sample using
TaqMan 2X Universal PCR Master Mix (Thermo Fisher Scientific, Grand Island, NY) using the manufacturer’s directions. MicroRNA expression levels were quantified using the
Applied Biosystems 7900HT Fast Sequence Detection System (Carlsbad, CA). Taqman
PRMT5 primers (product number: 4351370; assay ID: Hs01047356_m1) were used for
66 real-time PCR and GAPDH Taqman primers (product number: 4331182; assay ID:
Mm99999915_g1) were used as a normalization control (Thermo Fisher Scientific, Grand
Island, NY). ST7 primers were designed with the following sequences: TGA AAA TCA ACG
ACA ACT TG and ATA TTA GTG AGG AAG TGC CT and the FAM probe sequence CAC C A
AG T TC TAC GTG GCC CTAA. Cyclin D1 primers were designed with the following sequences, sense: CCGTCCATGCGGAAGATC and antisense: CCTCCTCCTCGCACTTCTGT and Power Sybr® Green (Applied Biosystems [Thermo Fisher Scientific, Grand Island,
NY]) dilution reagent was used for the real time PCR experiment. Expression levels were verified by three independent experiments.
Cell cycle
About 1 x 106 cells per condition were collected by centrifugation at 1200 rpm for 7 minutes. The supernatant was aspirated and the pellet was washed in 5 mL PBS pH 7.4
(Gibco, Grand Island, NY). The cells were again centrifuged for 7 minutes at 1200 rpm.
The resulting pellet was washed in sample buffer made with the PBS mentioned above with 0.1% glucose (Fisher Scientific, Pittsburgh, PA) added. The cells were again centrifuged using the same settings mentioned above. The supernatant was aspirated and the cell pellet was resuspended in 500 µL of sample buffer. While vortexing at a low to medium speed, 200 proof ethanol was slowly added dropwise to the cells to prevent precipitation. The cells were allowed to fix overnight for at least 16 hours at 4°C. The next day the cells were pelleted by centrifuging for 10 minutes at 2500 rpm. A master mix of staining solution was made for all the samples that included 89 µL of sample buffer, 1 µL
67
RNAse A (Qiagen, Valencia, CA) and 10 µL propidium iodide (BD Biosciences, San Jose,
CA) per sample. All samples were resuspended in 100 µL of the staining solution and incubated in the dark for 30 minutes at room temperature. After the 30 minute incubation, 400 µL of sample buffer was added to the cells and flow cytometry analysis was performed. The flow cytometry was performed on a FC500 Beckman Coulter flow cytometer (Beckman Coulter, Pasadena, CA).
Co-immunoprecipitation (Co-IP)
About 40 x 106 cells per condition were pelleted by centrifugation and rinsed once with pH 7.4 PBS. After the PBS wash the pellet was suspended in 500 µL of complete M-PER™ buffer (Thermo Fisher, Pittsburgh, PA) supplemented with 1X Halt™ protease and phosphatase inhibitors (Thermo Fisher, Pittsburgh, PA). The lysates were then precleared by adding 25 µL of well mixed agarose beads (Rockland, Limerick, PA) and incubating on a tube rotator for 30 minutes at 4°C. After 30 minutes the lysate was centrifuged for 3 minutes at 10,000 x g. The supernatant was transferred to a new tube and quantified for protein analysis using the Thermo Fisher BCA kit as described previously. Next, 2 tubes with 500 µg of protein were diluted in 500 µL of complete M-PER™ buffer as described above for each condition. 25 µL of the diluted protein was removed for input and combined with an equal volume of 2X sample loading buffer completed with β- mercaptoethanol and kept at -80°C until needed for western blotting. For the precipitation, 5 µg of the antibody of interest was added to 1 tube and 5 µg of IgG of the species of the antibody was added to the remaining tube. The tubes were allowed to
68 rotate overnight at 4°C. The next day 50 µL of the agarose beads used previously were added to both tubes. The tubes were allowed to rotate for 1 hour at 4°C. After the incubation, the tubes were centrifuged for 1 minute at 10,000 x g at 4°C. The supernatant was removed carefully to avoid loss of the beads and the beads were washed with 500
µL of M-PER™ buffer for three times with using the same centrifugation settings mentioned previously. After the last wash, the supernatant was removed and 100 µL of
2X sample loading buffer with β-mercaptoethanol was added to the beads. The tubes with the sample loading buffer and bead mixture and the input tubes were boiled for 10 minutes at 99°C. After boiling the tubes with beads were centrifuged at 10,000 x g for 3 minutes. Without disturbing the beads, 20 µL of input samples and IP sample were loaded onto a gel. For the western blotting procedure see previous description.
Chromatin immunoprecipitation (ChIP)
CHIP experiments were performed utilizing the MagnaChIP™ A Chromatin
Immunoprecipitation Kit (Millipore, Billerica, MA). Chromatin for each condition was extracted from 40 million cells. Sonication was performed with the Bioruptor® UCD-200
(Diagenode Inc, Denville, NJ). The chromatin was processed and prepared for real-time
PCR analysis per the kit instructions and with the reagents provided.
Nanostring
Total RNA for Nanostring analysis was extracted with Trizol reagent and washed and purified with buffers from MirVana. Nanostring profiling was performed with the
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Figure 3. PRMT5 protein is over-expressed in mantle cell lymphoma.
Protein isolated from primary MCL patient samples and MCL cell lines was analyzed for PRMT5 protein expression by western blot. Resting B cells were included as a negative control and Jeko is included as a positive control for the MCL patient blot. A) PRMT5 is overexpressed primary MCL patient samples numbered 1 through 16 and B) the MCL cell lines cc-MCL, Jeko, UPN-1, Granta, Mino, Rec-1, SP53 and Z138.
nCounter human v2 microRNA expression assay. Raw read data from the nCounter microRNA assay were analyzed by a biostatistician and converted to fold change values.
Statistical Analysis
Results were expressed as means ± standard deviation unless otherwise specified. F-tests for analysis of variance and paired t-tests were conducted in Microsoft Excel 2010 and used to generate p-values for comparisons between two groups and when multiple samples within different groups were used, respectively. Kaplan Meier analysis for engraftment models was performed in JMP Pro 12.
Results
PRMT5 is an important oncogenic target in mantle cell lymphoma
Dysregulation of PRMT5 has been shown to be a driver event in many malignancies
70
including lymphoma. In order to
characterize the role of PRMT5 in
MCL, the baseline expression of
PRMT5 protein was examined in
primary MCL patient samples and
commonly used MCL cell lines.
As expected, PRMT5 was
overexpressed in most primary
MCL patient samples (Figure 3A)
and in all MCL cell lines analyzed Figure 4. PRMT5 expression in MCL directly correlates with aggressive variants in mantle cell lymphoma. relative to normal resting B cells 53 MCL patients were grouped based on MCL variant as either classic, blastoid or pleomorphic and PRMT5 expression was (Figure 3B). In order to further examined in the samples by immunohistochemical analysis. Samples were either obtained from a malignant lymph node or investigate the differences of spleen. A) Immunohistochemical representations of PRMT5 expression in patient samples. The first panel demonstrates a positive PRMT5 stain that is mostly cytoplasmic while the second PRMT5 expression in primary panel shows a positive PRMT5 stain that is cytoplasmic and weakly nuclear. The third panel is a negative control. B) Graphical MCL patient samples, representation of the positivity and distribution of PRMT5 in MCL patient samples. Patient samples with the less aggressive classic variant of MCL mostly express PRMT5 in the cytoplasm and a few immunohistochemistry was samples are negative for PRMT5. As the MCL variant becomes more aggressive from blastoid to pleomorphic, PRMT5 positivity employed to detect PRMT5 increases and becomes more distributed into the nucleus. Additionally all of the samples in the highly aggressive pleomorphic variant were positive for PRMT5. cyt = cytoplasmic; expression in 53 MCL patient cyt/nuc = cytoplasmic and nuclear. samples were stratified into the variants: classic, blastoid or pleomorphic. Samples positive for PRMT5 were characterized as either containing solely cytoplasmic PRMT5 or cytoplasmic and weakly nuclear
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Table 3. Tabular representation of PRMT5 expression in MCL stratified by MCL variant.
PRMT5. Samples without any PRMT5 positivity were termed negative (Figure 4A). The less aggressive classic form of MCL demonstrated more cytoplasmic PRMT5 and several cases were negative for PRMT5, however the blastoid and pleomorphic variants of MCL displayed more nuclear PRMT5 and none of the highly aggressive pleomorphic MCL specimens were negative for PRMT5 (Figure 4B). Out of the 53 patient samples analyzed, only 11% were negative for PRMT5 (Table 3). The discovery that PRMT5 is expressed in the majority of samples along and that increased PRMT5 expression correlates with an increase in aggressiveness suggests that PRMT5 dysregulation is an important transforming event in MCL and one that could potentially be targeted with a therapeutic agent.
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Figure 5. Knockdown of PRMT5 results in a decrease of cell proliferation.
PRMT5 was knocked down with either siRNA or lenti-viral shRNA in Jeko MCL cell lines by treating the cells for 72 hours. A) Western blot analysis of PRMT5, cyclin D1 and sym-H4R3(2Me) in Jeko cells after PRMT5 knockdown show a moderate decrease in these proteins with either siRNA or lenti-viral shRNA treatment. B) Proliferation as measured by MTS assay after treatment with either si-PRMT5 or lenti-shPRMT5 show a decrease in proliferation with both siRNA and lenti-viral shRNA. Results are plotted relative to scramble control. *p<0.001.
Development of a PRMT5 small molecule inhibitor
Inhibition of PRMT5 with either small interfering RNA (siRNA) or shRNA delivered via lenti-virus in the MCL cell line Jeko led to a modest decrease in PRMT5 and its histone mark sym-H4R3(2Me) along with a slight decrease in cyclin D1 protein expression
(Figure 5A). Although the decrease in PRMT5 protein expression was slight with si- or shRNA knockdown there was a significant decrease in cellular proliferation indicating that these cells potentially require PRMT5 for normal cellular proliferation (Figure 5B).
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Figure 6. PRMT5 small molecule inhibitors are developed using an in silico model.
A) S-adenosyl-homocysteine (SAH) and arginine residue conformations docked to Human PRMT5 model (b) as compared to crystal structure of rat PRMT1 with co-crystallized SAH and substrate arginine (a). For clarity, protein molecular surface is generated omitting the residues covering the catalytic site face. Surface transparency is applied to show catalytic residues. Conserved catalytic residue interactions are reproduced in docking to hPRMT5 model as displayed. 10,000 compounds were screened for their ability to dock into this in silico model of human PRMT5. B) The top 8 compounds chosen for possessing the lowest binding energy to the in silicio model are shown.
Due to the inefficiency of PRMT5 knockdown in these cells by these methods, the possibility of creating a small molecule inhibitor of PRMT5 was explored. Since the molecular structure of human PRMT5 (hPRMT5) was unknown at the time, our organic chemist collaborators formed an in silico model of hPRMT5 using Modeller9v1 software.
The model was developed by exploiting the sequence homology of hPRMT5 and rat
PRMT1 (rPRMT1) and fitting the sequence of hPRMT5 into the structure of rPRMT1
(Figure 6A). The ChemBridge chemical library was then used to screen over 10,000
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Figure 7. Small molecule inhibitors of PRMT5 inhibit the proliferation of MCL cells.
MTS assay was conducted using the MCL cell lines Jeko and Mino treated with various concentrations of the 8 candidate PRMT5 inhibitors. Cells were treated for 24 and 48 hours with the compounds prior to addition of the MTS reagent. Results are plotted relative to untreated (UT) control. CMP3 and CMP5 showed the best inhibition of proliferation in these cells.
compounds for their ability to bind into the SAM docking site and block the methyltransferase activity of PRMT5 with the lowest energy expenditure. Eight candidate compounds were chosen based on their low binding energy to the desired site (Figure
6B). The eight candidate compounds were subjected to rigorous testing in order to determine their ability to inhibit the proliferation of malignant MCL cells as well as their ability to specifically inhibit the methyltransferase activity of PRMT5 without affecting the methyltransferase activity of other PRMTs. Compound 3 (CMP3) and compound 5
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Figure 8. CMP3 and CMP5 selectively knockdown of symmetric-H4R3(2Me) in the MCL cell lines Jeko and Mino.
Confocal microscopy was utilized to investigate the ability of the compounds to block the methyltransferase activity of PRMT5 by examining the expression of the histone mark of PRMT5 symmetric-H4R3(2Me). The PRMT1 specific mark asymmetric-H4R3(2Me) was included as a negative control. Jeko and Mino MCL cell lines were treated with each compound for 48 hours prior to processing for confocal. Green = asym or sym- H4R3(2Me) and blue = nuclear DAPI stain. Both CMP3 and CMP5 inhibit only symmetric-H4R3(2Me).
(CMP5) inhibited cell growth in the MCL cell lines Jeko and Mino better than the other six compounds as measured by the MTS proliferation assay (Figure 7).
Immunofluorescence analysis of MCL cell lines after treatment with the candidate compounds also determined that CMP3 and CMP5 treatment resulted in the loss of the
PRMT5 histone mark sym-H4R3(2me) without affecting the PRMT1 histone mark asym-
H4R3(2me) (Figure 8). CMP3 and CMP5 were able to inhibit the methyltransferase activity of PRMT5 as measured by a fluorometric
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Figure 9. Ability of candidate compounds to block PRMT5 methyltransferase activity.
A) The ability of the 8 candidate compounds to block methyltransferase activity was measured by fluorometric methyltransferase assay. All compounds were used at a concentration of 100 µM and methylation activity was plotted relative to the DMSO control. Only CMP3 and CMP5 significantly inhibit the methyltransferase activity of PRMT5. Since CMP5 appeared to inhibit methyltransferase activity the most it was used for subsequent experiments. B) The fluorometric assay described previously was used to determine that neither CMP3 nor CMP5 inhibited the methyltransferase activity of PRMT1. C) Various concentrations of CMP5 were used to test the ability of the compound to block the methyltransferase activity of PRMT5. Methyltransferase activity was plotted relative to untreated (0µM) control. CMP5 was able to inhibit over 50% of the methyltransferase activity of PRMT5 at 100 µM.
methyltransferase assay (Figure 9A). Additionally neither CMP3 nor CMP5 inhibited the methyltransferase activity of PRMT1 indicating that both compounds specifically blocked the methyltransferase activity of PRMT5 only (Figure 9B). Since CMP5 inhibited the methyltransferase activity of PRMT5 to a greater existent than CMP3 it was used for all subsequent experiments. Concentration dependent analysis of CMP5 demonstrated it was able to inhibit over 50% of the methyltransferase activity of PRMT5 at 100 µM
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Figure 10. CMP5 selectively inhibits the methyltransferase activity of PRMT5.
The ability of CMP5 to inhibit the methyltransferase activity of PRMT5 was tested with pure histones. PRMT4, a type I PRMT, and PRMT7, another type II PRMT were included as controls. CMP6 was included as a non- inhibiting control. All drugs were used at a concentration of 100 µM. A) In the presence of total histones CMP5 only inhibits the methyltransferase activity of PRMT5. Additionally in the presence of B) histone H4 or C) histone H3 CMP5 only inhibits the methyltransferase activity of PRMT5.
(Figure 9C). To further confirm the specificity of CMP5 for PRMT5, the ability of the compound to inhibit the methyltransferase activity of PRMT5 was tested along with its effects on PRMT4, a type I methyltransferase, or PRMT7, another type II methyltransferase. CMP5 only inhibited PRMT5 in the presence of total histones (Figure
10A), or histone H4 (Figure 10B) and histone H3 alone (Figure 10C) and demonstrated no ability to inhibit the methyltransferase activity of PRMT4 or 7. These results
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Figure 11. Inhibition of PRMT5 activity with CMP5 is only toxic in malignant MCL cells.
Resting B cells from normal donors, PBMCs from MCL primary patients and MCL cell lines were treated with various concentrations of CMP5. Cell viability was measured by annexin/PI flow cytometry and results are plotted relative to DMSO or untreated (0) control. A) The viability of B cells unaffected by CMP5 treatment up to 100 µM. Results are pooled from 2 separate donors. B) CMP5 is toxic to MCL patient samples and viability is reduced around 50 µM at 48 hours in these samples. Results are pooled from 3 MCL patient samples. C) CMP5 is toxic to MCL cell lines. Results are pooled from 3 MCL cell lines (Jeko, Mino and Z138). *p<0.01; **p<0.001; ***p<0.0001
established that CMP5 specifically blocked the methyltransferase activity of PRMT5 without affecting the methyltransferase activity of other type II PRMTs, such as PRMT7 or
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the methyltransferase activity of
type I PRMTs such as PRMT4.
CMP6 was included as a negative
control as it was previously
Figure 12. CMP5 treatment affects the histone mark determined to have no effect on symmetric-H4R3 (2Me) in MCL. the methyltransferase activity of Jeko and Mino cells were treated with CMP5 for 24 hours. CMP5 treatment resulted in loss of the PRMT5 histone mark (middle band) but did not affect the protein expression of PRMT5. PRMT5. Positive control (+ctrl) = Jeko cells treated with 10 nM bortezomib. CMP5 treatment decrease cell
viability and proliferation of
MCL cell lines and primary patient samples
In order to determine the effect of CMP5 on normal B cells versus malignant samples, resting B cells, MCL patient samples and MCL cell lines were treated with various concentrations of CMP5 for 24, 48 and 72 hours and the viability of each group was determined by annexin/PI flow cytometry. Resting B cells were not significantly sensitive to CMP5 treatment up to 72 hours at 100 µM (Figure 11A). In contrast both MCL patient samples and cell lines were sensitive to CMP5 treatment and displayed a drastic decrease in cell viability relative to untreated cells (Figure 11B and Figure 11C). Additionally, treatment with CMP5 resulted in a decrease in the protein expression of the PRMT5 histone mark sym-H4R3 (2me) without causing a change in the protein expression of
PRMT5 itself indicating that the compound only regulates the methyltransferase activity of PRMT5 and not the expression of PRMT5 (Figure 12). In accordance with the
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Figure 13. CMP5 treatment results in the activation of apoptotic pathways in the MCL cell lines Jeko and Mino.
Jeko and Mino cells were treated for 24 hours with the indicated concentration of CMP5 A) Caspase-3 cleavage (lower band) in only occurs in Jeko cells with CMP5 treatment. B) Parp is also cleaved (lower band) in Jeko cells with CMP5 treatment. C) There is an increase in the pro-apoptotic (lower band) product of MCL- 1 in Jeko and Mino cells with CMP5 treatment. Positive control (+ctrl) = Jeko cells treated with 10 nM bortezomib. apoptosis data, Jeko cells treated with CMP5 induced both caspase-3 (Figure 13A) and
PARP (Figure 13C) cleavage indicating inhibition of PRMT5 in these cells results in activation of the extrinsic apoptotic and DNA damage-repair pathways respectively.
Interestingly CMP5 treatment in Mino cells does not result in either caspase-3 or PARP cleavage however there is an increase in the pro-apoptotic short form of the Bcl-2 member MCL-1 in both Jeko and Mino cells (Figure 13B). These results indicate that potentially PRMT5 promotes malignancy though different pathways in these two cells lines and as a result, inhibition of PRMT5 activity activates different apoptotic pathways in these two cell lines.
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Figure 14. PRMT5 inhibition with CMP5 treatment in MCL attenuates cyclin D1 expression.
Jeko and Mino cells were treated for 24 hours with the indicated concentrations of CMP5 and lysates were analyzed by western blot. A) Western blot analysis of Bcl-2 family proteins indicate there is no change in their expression with CMP5 treatment. B) There is also no change in AKT or C) ERK family proteins with CMP5 treatment in MCL. D) Cyclin D1 expression is loss in MCL cell lines after treatment with CMP5. The positive control (+ ctrl) is Jeko cells treated with 10 nM Bortezomib.
PRMT5 inhibition with CMP5 affects the cell cycle pathway in MCL
CMP5 was originally designed and employed to be used as a tool to elucidate the pathways controlled by PRMT5 to drive malignancy in MCL with the potential to be used as a therapeutic in the future. In order to glean the role of PRMT5 in MCL, the effect of
CMP5 treatment on pathways commonly known to be dsyregulated in MCL was examined via western blotting. The pro-survival BCL-2 proteins are commonly overexpressed in MCL in up to 97% of cases and may be responsible for mediating apoptotic-resistance in this disease [205]. However, treatment with CMP5 does not affect the expression of the pro-survival BCL-2 proteins BCL-2 and BCL-XL in Jeko or Mino cells
(Figure 14A). CMP5 treatment also does not affect the expression of the pro-apoptotic
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Figure 15. Inhibition of PRMT5 activity in MCL results in time dependent loss of cyclin D1.
A) MCL cell lines treated with CMP5 for 24 hours show a decrease in cyclin D1 protein levels. Time course dependent treatment of Jeko (B) and Mino (C) MCL cell lines results in the loss of cyclin D1 protein by 3 hours with CMP5 treatment as demonstrated by western blot analysis. CMP5 treatment concentrations FC- mMCL = 25 µM, UPN-1 = 30 µM, Jeko and Z138 =50 µM, Granta, and Mino = 60 µM and Rec-1 and SP53 = 75 µM.
BCL-2 protein BAX. The lack of change in Bax expression again suggests that the apoptosis subsequent to CMP5 treatment mainly occurs as a result of activation of the extrinsic apoptotic pathway mediated by caspase proteins in Jeko cells rather than the intrinsic pathway which is mediated by BCL-2 proteins. The AKT pathway is also constitutively overexpressed in MCL and contributes to maintaining cyclin D1 expression and cell cycle dysregulation[223]. CMP5 treatment does not affect the protein expression of either Akt or phosphorylated Akt in Jeko or Mino cells after 24 hours (Figure 14B).
The ERK 1/2 pathway is also amplified in MCL and PRMT5 has been shown to play a role in mediating the ERK 1/2 signaling pathway thus we examined ERK expression in MCL with CMP5 treatment [224,225]. Blockade of PRMT5 with CMP5 activity does not cause changes in Erk or phosphorylated ERK protein (Figure 14C), however, CMP5 treatment
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Figure 16. Inhibition of PRMT5 with CMP5 affects the cell cycle in MCL.
A) Treatment of the MCL cell lines Jeko and Mino with CMP5 for 24 hours results in protein loss of cyclin D members and their binding partner CDK4 as determined by western blot analysis. There is no change in cdk6 expression with CMP5 treatment. B) CMP5 treatment does not change the expression levels of CCND1 mRNA as determined by real-time PCR.
does result in a significant decrease in cyclin D1 protein in Jeko and Mino cells (Figure
14D). Additionally the loss of cyclin D1 protein with CMP5 treatment was seen in all MCL cell lines tested (Figure 15A). Time course analysis revealed that cyclin D1 protein expression was completely demolished within 3 hours of CMP5 treatment in both Jeko and Mino cells (Figure 15B and Figure 15C). We also examined at the protein expression of cyclin D2 and D3 since studies have shown that the loss of cyclin D1 protein can be compensated for by the upregulation of the other cyclin D proteins, cyclin
D2 and/or cyclin D3 [206,226]. CMP5 treatment also caused a loss of either cyclin D2 in
Mino cells or cyclin D3 in Jeko cells after 24 hours (Figure 16A). Moreover, the main cyclin D binding partner, CDK4, was also loss with CMP5 treatment, while the alternate cyclin D binding partner, CDK6, was unaffected. Interestingly, there was no change in the expression level of cyclin D1 mRNA (Figure 16B) insinuating that the change in cyclin D1 protein expression could be attributed to a post-translational modification. Subsequent
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Figure 17. Inhibition of PRMT5 activity with CMP5 causes G2/m cell cycle arrest.
Jeko A) and Mino B) cells were treated with CMP5 for 24 hours at the indicated concentrations and subjected to cell cycle analysis via flow cytometry. CMP5 treatment resulted in G2/M arrest.
cell cycle analysis revealed that treatment with CMP5 resulted in G2/M arrest in both
Jeko and Mino cells (Figure 17A and Figure 17B). Cyclin D1 binds to CDK4 to form an activated complex that can phosphorylate and inactivate Rb protein. Rb normally binds to and inhibits E2F transcription factors preventing cell cycle progression; however phosphorylated Rb is unable to bind to and inhibit the EF2 transcriptional machinery which allows for the production of the E2F proteins and cyclin E required for progression to the S-phase of the cell cycle. Since CMP5 treatment decreased the protein expression of cyclin D proteins and their binding partner CDK4 we decided it was reasonable to examine the expression of Rb and downstream E2F proteins. Not surprisingly, blockade of PRMT5 methyltransferase activity with CMP5 led to a decrease in phosphorylated Rb protein expression (Figure 18). In Jeko cells CMP5 treatment had no effect on the expression of total Rb however the band for total Rb in the treated conditions is slightly
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lower, potentially denoting loss of
a post-translational modification, in
this case phosphorylation. In Mino
cells CMP5 treatment resulted in a
decrease in total Rb protein though
to a lesser extent than
Figure 18. CMP5 treatment causes changes in Rb phosphorylated Rb. The Rb band in expression in MCL cell lines.
Jeko and Mino cell lines were treated for 24 hours with CMP5 Mino cells is also lower again and the expression of Rb and 3 different phospho-Rb proteins was analyzed via western blot. CMP5 treatment resulted in a suggesting the loss of loss of the phosphorylated forms of Rb concurrent with a slight decrease in the molecular weight of total Rb. phosphorylation. CMP5 also resulted in a loss of E2F1 and 2 proteins in both Jeko and Mino (Figure 19A). Cyclin E regulates the G1 to S progression of the cell cycle and is also an E2F gene product.
Ablation of PRMT5 methylation activity with CMP5 was also able to decrease the protein
(Figure 19B) and mRNA expression of cyclin E (Figure 19C) and, to a lesser extent, the protein expression of CDK2 (Figure 19B) the binding partner of cyclin E. Western blot analysis of cyclin A and cyclin B1, which control the progression from S to G2 and from
G2 to M progression respectively, after CMP5 treatment revealed little to no effect on these proteins (Figure 20). Interestingly, cdc2 (CDK1), the binding partner for cyclin B1, decreased with treatment. Taken together these data indicate that PRMT5 plays a role in driving cell cycle progression in MCL, particularly by controlling or cooperating with the cyclin D pathway, and that attenuation of the methyltransferase activity of PRMT5 results
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Figure 19. PRMT5 inhibition with CMP5 results in a loss of E2F target gene and protein products.
Jeko and Mino cells were treated with CMP5 for 24 hours with the indicated concentrations. Protein was collected for western blot analysis and RNA was collected for mRNA analysis by quantitative real time PCR. A) E2F1 and E2F2 protein expression is loss in MCL with CMP5 treatment. B) CMP5 treatment also reduced cyclin E protein expression. Cdk2 expression is modestly reduced in Jeko cells with treatment. C) CCNE (cyclin E) mRNA is significantly reduced with CMP5 treatment in Jeko and Mino cells. Graph is plotted as a ratio. Compared to the DMSO solvent control *p<0.05.
in loss of cell cycle progression, proliferation and an increase in apoptosis of the cells.
PRMT5 and cyclin D1 associate in MCL
Since the expression of cyclin D1 protein was sensitive to inhibition of the methyltransferase activity of PRMT5, we wanted to determine if PRMT5 and cyclin D1 directly interacted with each other in MCL. Immunoprecipitation of PRMT5 in Jeko cells revealed that cyclin D1 co-precipitated with PRMT5 (Figure 21). These experiments verify that cyclin D1 and PRMT5 associate with each other and potentially exert control over each other.
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Inhibition of PRMT5 activity
does not increase the activity of
the proteasome
Cyclin D1 is phosphorylated by
GSK3β which targets it for
Figure 20. CMP5 treatment does not affect the expression ubiquitination and subsequent of other cyclins but decreases the expression of cdc2. degradation by the proteosome. In The MCL cell lines Jeko and Mino were treated with CMP5 for 24 hours at the indicated concentrations and protein was collected for western blot analysis. CMP5 treatment did not order to determine if the loss of affect the protein expression of cyclin A or B1 but it did decrease the expression of the cyclin B1 binding partner cdc2. cyclin D1 protein expression with
inhibition of PRMT5 activity was due to increased proteasomal degradation of cyclin D1, the protein expression of phosphorylated cyclin D1 and GSK3β was examined. Treatment with CMP5 in Jeko and
Mino cells decreased phosphorylated cyclin D1, and in Jeko cells GSK3β protein expression was decreased with treatment while in Mino cells GSK3β protein expression was increased (Figure 22). These results were inconclusive therefore Jeko and Mino cells were treated with the proteasomal inhibitor bortezomib for various time conditions.
Treatment with bortezomib for 1 hour with a pre-established dose [227] partially rescued
Jeko cells treated with 75 µM CMP5 from apoptosis (Figure 23A first panel). Bortezomib treatment also modestly rescued Mino cells treated with 75 µM CMP5 from apoptosis but not cells treated with 100 µM (Figure 23A second panel). In Jeko cells cyclin D1 expression was modestly rescued with 30 minute and 1 hour bortezomib treatment
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however the effect was lost by 2 hours (Figure 23B).
Thus, although bortezomib is initially able to rescue
apoptosis and cyclin D1 protein expression, the
effect is not sustained. This implies that there is Figure 21. PRMT5 and cyclin D1 associate in MCL. another mechanism responsible for the loss of cyclin Jeko cells were immunoprecipitated with PRMT5 antibody, and probed for cyclin D1 protein expression that is proteasome- D1 and HDAC2 on a western blot. Cyclin D1 was able to co-immunoprecipitate with PRMT5 in Jeko cells. HDAC2, a independent. known binding partner of PRMT5 in the SWI/SNF complex was included as an Inhibition of PRMT5 results in a loss of c-myc immunoprecipitation control.
The transcription factor c-myc plays a role in cellular transformation by regulating apoptotic and cell cycle pathways. Although rare, small subsets of MCL patients over-express myc possibly due to either a translocation or gene amplification event, and typically present with more aggressive variants of MCL [228].
Additionally murine models that overexpress c-myc in conjunction with mutations to the
INK4A or p53 pathways develop MCL-like lymphomas [229]. Treatment of Jeko and Mino cells with CMP5 caused a decrease in the protein expression of c-myc (Figure 24A).
Similar to the results with cyclin D1, loss of c-myc was time dependent and occurred quickly; within 6 hours for Jeko and within 3 hours for Mino. Surprisingly, real-time PCR analysis in Jeko cells revealed that c-myc mRNA increased after CMP5 treatment (Figure
24B). These data demonstrated that similar to that which occurred with cyclin D1, PRMT5 likely acts in a post-translational manner to control the protein expression of c-myc, which explains the lack of correlation between the mRNA and protein expression with
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CMP5 treatment. Blockade
of the methyltransferase
activity of PRMT5
decreases the protein
expression of c-myc and
potentially activates Figure 22. CMP5 treatment does not induce the proteasomal degradation pathway for cyclin D1. compensatory Jeko and Mino cells were treated with CMP5 for 24 hours at the indicated concentrations, and probed for phosphorylation cyclin D1 and mechanisms that result in GSK3β on a western blot. CMP5 treatment decreased phosphorylated cyclin D1. GSK3β expression was decreased in Jeko cells but increased in an increase in mRNA Mino cells with CMP5 treatment.
expression.
Inhibition of PRMT5 results in a de-repression of tumor suppressors
PRMT5 has been shown repress a number of tumor suppressors, including suppressor of tumorigenicity 7 (ST7), by symmetrically dimethylating histone proteins in their promoter region [125]. Based on this knowledge, we sought to determine what effect blockade of the methyltransferase activity of PRMT5 had on the expression of ST7. Confocal microscopy analysis proved that Jeko and UPN-1 cells treated with CMP5 for 48 hours saw an increase in the expression of ST7 protein (Figure 25A). Additionally, quantitative real-time PCR analysis demonstrated an increase in ST7 mRNA in the MCL cell lines Jeko,
Mino and UPN-1 treated with CMP5 (Figure 25B). Furthermore, chromatin immunoprecipitation (ChIP) of PRMT5 revealed a significant loss of PRMT5 recruitment to the promoter of ST7 when Mino cells were treated with CMP5 (Figure 25C). Jeko cells
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Figure 23. Proteasome inhibition does not rescue MCL cells from the effects of CMP5.
Jeko and Mino cells were pre-treated with 10 nM bortezomib (PS-341) for 4 hours prior to treatment with CMP5 at the indicated doses. A) Viability was assessed by annexin V/PI flow cytometry. Bortezomib cannot rescue MCL cells from the toxic effects of CMP5. B) Western blot of cyclin D1 in Jeko cells shows that pretreatment with bortezomib partially rescued cyclin D1 expression 30 minutes after CMP5 treatment however the rescue was not sustained pass 2 hours.
also showed a decrease in PRMT5 recruitment to ST7 with CMP5 although it did not reach statistical significance. We also examined the expression of the tumor suppressor
C/EBPβ with and without CMP5 treatment in MCL. C/EBPβ has been shown to inhibit cell growth and modulate the immune response in various malignancies and C/EBPβ target genes can be inhibited by the methyltransferase activity of PRMT5 [230,231]. CMP5
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Figure 24. Inhibition of PRMT5 with CMP5 results in a loss of c-myc protein expression in Jeko and Mino cells.
A) Western blot analysis revealed that Jeko and Mino cells treated with CMP5 lose c-myc protein expression. Jeko cells were treated with 75 µM CMP5 and c-myc protein expression began to fall around 6 hours post treatment. Mino cells were treated with 100 µM CMP5 and c-myc protein expression began to fall around 3 hours post treatment. B) Jeko cells treated with CMP5 do not show a decrease in c-myc mRNA as measured by real time PCR. *p<0.001.
treatment resulted in a significant induction of C/EBPβ protein and mRNA in both Jeko and Mino cells (Figure 26A and Figure 26B). Additionally, similar to ST7, ChIP analysis indicated a significant loss of PRMT5 recruitment to the promoter of C/EBPβ in both Jeko and Mino cells with CMP5 treatment (Figure 26C).
Since we saw an attenuation of c-myc expression with PRMT5 methyltransferase inhibition via CMP5, we decided to examine the expression of the tumor suppressor programmed cell death 4 (PDCD4), which specifically inhibits c-myc induced cellular proliferation and survival [232]. Not surprisingly, treatment of Jeko cells with CMP5 resulted in a significant increase in PDCD4 protein and mRNA expression (Figure 27A and Figure 27B). Furthermore, both Jeko and Mino cells demonstrated an upregulation in PDCD4 expression as early as 3 hours post-CMP5 treatment (Figure 27C).
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Figure 25. Inhibition of PRMT5 methyltransferase activity with CMP5 results in the de-repression of ST7.
A) CMP5 treatment for 48 hours in Jeko (75 µM) and UPN-1 (30 µM) MCL cell lines resulted in an increase in the expression ST7 and a decrease in the expression of PRMT5 epigenetic mark symmetric dimethyl-H4R3 as measured by confocal microscopy. The first picture in each 4 panel group (top left) represents ST7 (green), the second picture (top right) represents a phase contrast image of the cells, the third picture (bottom left) denotes the nuclear stain DAPI (blue), and the fourth picture is an overlay of the ST7 and DAPI stains. B) CMP5 treatment for 48 hours in Jeko (75 µM), Mino (100 µM) and UPN-1 (30 µM) MCL cell lines resulted in an increase in the mRNA expression of ST7. C) Chromatin immunoprecipitation of PRMT5 demonstrated a loss of PRMT5 recruitment to the ST7 promoter with CMP5 treatment in Mino cells. *p<0.05. The results were not significant in Jeko cells.
These experiments indicate that PRMT5 is responsible for perpetuating transformation and malignancy in MCL in part by suppressing the expression of key tumor suppressors.
Inhibition of the methyltransferase activity of PRMT5 with a small molecule inhibitor, successfully de-repressed the expression of ST7, C/EBPβ, and PDCD4 which likely was at
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Figure 26. Treatment with CMP5 in MCL cell lines increases the expression of C/EBPβ.
A) Western blot results demonstrated that inhibition of the methyltransferase activity of PRMT5 with 24 hour CMP5 treatment resulted in an increase in the protein expression of C/EBPβ in the MCL cell lines Jeko and Mino. B) Real-time PCR analysis demonstrated an increase of C/EBPβ mRNA in MCL cell lines treated with CMP5. *p-value<.05; **p-value<0.01 C) Chromatin immunoprecipitation experiments determined that PRMT5 recruitment to the promoter of C/EBPβ was loss with CMP5 treatment. *p-value<0.05.
least partially responsible for the decrease in cell proliferation and the increase in apoptosis in MCL cell lines treated with the LC50 dose.
Inhibition of PRMT5 perturbs microRNA networks in MCL
On account of the rapid loss of protein expression of both cyclin D1 and c-myc, we hypothesized that inhibition of PRMT5 potentially perturbed miR networks in MCL. In order to test our hypothesis we submitted total RNA from 5 MCL cell lines treated with
CMP5 for 72 hours and 1 MCL cell line transduced with shPRMT5 for Nanostring
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Figure 27. Inhibition of PRMT5 methyltransferase activity with CMP5 results in an increase in the expression of the tumor suppressor PDCD4.
A) The MCL cell line Jeko was treated with CMP5 at the indicated concentrations for the indicated timepoints. PDCD4 protein expression is increased with CMP5 relative to the DMSO treated control as analyzed by western blot. B) Quantitative real-time PCR demonstrated that PDCD4 mRNA expression is increased in Jeko cells treated with CMP5 at the indicated timepoints for the indicated concentrations with the most robust increase at 50 µM for 48 hours. C) Western blot analysis revealed that PDCD4 protein expression increased within 3 hours post CMP5 treatment in both Jeko (50 µM) and Mino (75 µM) MCL cell lines.
nCounter human microRNA expression analysis. The Jeko, Mino, SP53, UPN-1 and cc-
MCL cell lines were treated with CMP5 and in addition the Jeko cell line was transduced with shPRMT5. Solvent controls, in this case DMSO, or scramble treated cells were also analyzed simultaneously. Changes in miR expression were deemed significant if there was a 2 fold or more increase or decrease relative to the control samples. To compile a list of potential candidate miRs we only considered miRs whose expression changed significantly in 3 or more of the cell lines. Based on these criteria, Nanostring analysis
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Table 4. List of microRNAs that increase at least 2 fold in 3 or more MCL cell lines when treated with CMP5 or shRNA. revealed 17 miRs that were upregulated (Table 4) and 18 miRs that were downregulated
(Table 5) with PRMT5 knockdown. The majority of the upregulated miRs are novel miRs that have few or no known validated gene targets. In contrast most of the downregulated miRs are known oncomiRs that promote the translation of cell cycle and other oncoproteins.
Treatment with CMP5 has a synergistic effect when used in conjunction with other anti-tumor drugs
The typical treatment regime for malignancies such as MCL involves the simultaneous use of multiple chemotherapeutic agents in order to increase the effectiveness of the treatment and progression-free survival of patients. Since we are interested in epigenetics, we tested CMP5 in conjunction with other drugs that target epigenetic
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Table 5. List of microRNAs that decrease at least 2 fold in 3 or more MCL cell lines when treated with CMP5 or shRNA along with common genes that they target.
modifiers. Trichostatin A (TSA) inhibits class I and II histone deacetylases (HDACs), a group of epigenetic modifiers that removes acetyl groups from histone proteins in order
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Figure 28. The PRMT5 inhibitor CMP5 acts synergistically with other anti-tumor drugs used to treat MCL.
The MCL cell lines Mino and Jeko were treated with sub-lethal doses (LC25) of the DNA methyltransferase inhibitor 5-azacitidine (5-Aza = 500 nM) and the HDAC I and II inhibitor trichostatin A (TSA=50 nM) in addition to CMP5 (25 µM). A DMSO treatment group was included as a negative control. A) Viability was assessed via annexin V and PI flow cytometry. The addition of CMP5 to either TSA or 5-AZA resulted in a decrease in cell viability. The combination of TSA, 5-AZA and CMP5 (combo) resulted in the greatest decrease in cell viability as compared to the other treatment regimens. *p<0.05; **p<0.001 B) Low dose combination TSA, 5-AZA and CMP5 treatment resulted in a greater loss of cyclin D1 and symmetric dimethyl-H4R3 protein expression as compared to either single agent or double agent treatment as measured by western blot.
to silence genes. 5-Azacytidine (5-AZA) inhibits DNA methyltransferases (DNMTs), a
98 group of epigenetic modifiers that silences gene expression by transferring methyl- groups directly onto the DNA molecular itself. Both HDACs and DNMTs are known to form oncogenic complexes with PRMT5 to silence critical tumor suppressor genes in malignancies [190,233]. In order to assess the efficacy of combination TSA, 5-AZA and
CMP5 treatment in MCL, Jeko and Mino cells were treated with the LC25 dose of each of the drugs either alone or in combination. Neither of the drugs alone resulted in a significant decrease in cell viability, however, either 5-AZA or TSA in combination with
CMP5 caused a significant decrease in viability albeit not lower than 50% (Figure 28A).
Use of all three agents resulted in a dramatic decrease in cell viability and cyclin D1 and symmetric dimethyl-H4R3 protein expression in both Jeko and Mino cells (Figure 28B).
These encouraging results indicate that PRMT5 could potentially be used in conjunction with other chemotherapeutic drugs to cause a dramatic attenuation of malignant cellular growth.
Discussion
The epigenetic regulator PRMT5 has been implicated as an oncogene in a number of malignancies, thus, unsurprisingly, all of the MCL cell lines surveyed in this study overexpressed PRMT5 protein as compared to normal resting B-cells. Additionally, a significant portion of MCL patient samples expressed PRMT5 in a grade-dependent manner. Immunohistochemical analysis demonstrated that MCL patients with the less aggressive and slower progressing classic variant were either negative for PRMT5 expression or expressed cytoplasmic PRMT5, however, as MCL grade progressed to more
99 aggressive blastoid and pleomorphic variants, the incidence of PRMT5 positivity increased concomitant with nuclear expression of PRMT5. More nuclear expression of
PRMT5 allows it to achieve greater contact with the genetic machinery of the cell and promote transformation and malignancy by silencing key regulatory genes via its methyltransferase activity. It is unclear why the one pleomorphic patient included in the western blot failed to express PRMT5 protein, however it is possible the patient had a low white blood cell count at the time of procurement therefore not enough PRMT5 positive malignant cells were present in the sample to be able to be detected by western blot. The fact that only 11% of the 53 examined samples were negative for PRMT5 coupled with the increased incidence of nuclear PRMT5 as the disease becomes more aggressive suggests that PRMT5 plays an integral role in the establishment and maintenance of this malignancy.
In order to explore further the role of PRMT5 in MCL we collaborated with the department of Pharmacy here at The Ohio State University to design and synthesize small molecule inhibitors of PRMT5. Our collaborators designed an in silico model of hPRMT5 by fitting the sequence of hPRMT5 to the structure of rPRMT5, a model that was proven to be remarkably close to the actual molecular structure of PRMT5 that was published years later [128]. Using the Chembridge library of compounds we were able to find a compound, CMP5, which selectively docked into the SAM cofactor binding pocket on PRMT5 with a low binding energy, effectively preventing the methyltransferase activity of PRMT5. CMP5 treatment of MCL cell lines resulted in a decrease in the protein
100 expression of the PRMT5 epigenetic mark symmetric H4R3 but had no effect on the
PRMT1 epigenetic mark asymmetric H4R3. Additionally, CMP5 did not affect the methyltransferase activity of the type I PRMT1 or the other type II PRMT7, indicating that our compound specifically inhibited the methyltransferase activity of PRMT5 only.
Treatment of MCL primary patient samples and cell lines with CMP5 also resulted in decreased proliferation, and increased apoptosis while there was no effect on normal resting B-cells that express little to no PRMT5 protein. The increase in apoptosis was concomitant with an increase in the protein expression of several apoptotic regulators such as cleaved PARP and caspase 3 as well as the small apoptotic isoform of MCL-1.
These data suggest that PRMT5 plays a critical role in promoting transformation and survival in MCL.
There are several dysregulated pathways in MCL including the BCL-2, cell cycle, and
PI3K/AKT/MTOR pathways, however of particular interest, blockade of PRMT5 methyltransferase activity only affected the cell cycle pathway but was enough to cause significant cell death in MCL cell lines. Notably, CMP5 treatment reduced the protein expression of cyclin D1, its binding partner CDK4 as well as the other cyclin D members cyclin D2 and D3. Intriguingly, cyclin D1 protein expression fell quickly and did not correlate with mRNA expression indicating that PRMT5 may control cyclin D1 post- translationally. Potentially methylation of one or more arginine residues on cyclin D1 by
PRMT5 helps to stabilize cyclin D1 similar to that which occurs with p53 [234].
Furthermore, the loss of cyclin D1 protein expression lead to subsequent G2M cell cycle
101 arrest as opposed to the G1 arrest anticipated with cyclin D1 loss. Cyclin D1 is typically elevated in G1 phase to promote mRNA and protein synthesis required for the upcoming cell division. During S-phase cyclin D1 is downregulated to allow for DNA synthesis but is subsequently upregulated again during the G2/M portion of the cell cycle in order to promote cell division and continued proliferation for the next cycle of cell division. Under normal physiological conditions, cyclin D1 protein elevation during the G2/M phase occurs in a post-translational, Ras-dependent, mRNA independent manner [235]. Rather, the increase in cyclin D1 protein in the G2/M phase is a consequence of eI4FE mediated shuttling of cyclin D1 from the nucleus to the cytoplasm. Additionally, efficient translation of cyclin D1 occurs by an eIF4E-mediated 5’-cap dependent mechanism; a process that requires PRMT5 to facilitate the interaction between eIF4E and target mRNA
[236]. Thus in MCL perhaps PRMT5 is required for the efficient translation and shuttling of cyclin D1 protein during the G2/M phase of the cell cycle and, inhibition of PRMT5 has immediate detrimental effects on this pathway, resulting in G2/M cell cycle arrest and apoptosis.
E2F genes are required for the cell cycle to proceed from the G1 to the S-phase, however, Rb can bind to the E2F promoter and prevent transcription of these genes. To combat this, the cyclin D1/CDK4 complex phosphorylates Rb, releasing it from the E2F promoter and allowing E2F genes, such as cyclin E, to be transcribed. Treatment of MCL cell lines with the PRMT5 inhibitor ablated the protein expression of phosphorylated Rb with little to no change in the protein expression of total Rb, aside from a slight change
102 in molecular weight. The slight change in molecular weight of the Rb protein in cell lysates treated with the PRMT5-inhibitor CMP5 correlated with loss of a post- translational modification, in this case phosphorylation. Moreover, we were able to demonstrate that loss of Rb phosphorylation restored the inhibitory effects of Rb as can be seen with the decrease in the protein expression of E2F1, E2F2 and cyclin E. CMP5 treatment also resulted in the loss of the expression of CDK2, the binding partner of cyclin E. Overall these results imply that PRMT5 is not only necessary for the growth of
MCL but it is necessary to perpetuate the dysregulated growth program in MCL and allow cells to continue to proliferate.
Based on the results with the PRMT5 inhibitor, CMP5, and the loss of cyclin D1 we theorized that PRMT5 directly interacted with cyclin D1 in MCL and by some means prevented its proteasomal degradation. While immunoprecipitation experiments confirmed that PRMT5 and cyclin D1 directly interacted with each other, blockage of the proteasome with bortezomib (PS-341) was unable to reverse the loss of cyclin D1 protein with CMP5 treatment implying that PRMT5 is relevant to the cyclin D1 pathway in MCL by some other means, most likely via post-translational modification as mentioned previously. Moreover, we were unable to find a correlation between the expression of
GSK3β, a protein that phosphorylates cyclin D1 and targets it for degradation, and CMP5 treatment, further indicating that PRMT5 has no effect on the proteasomal degradation of cyclin D1.
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Similar to that which occurred with cyclin D1, CMP5 treatment resulted in a decrease in c-myc protein expression. Also similar to cyclin D1, c-myc loss with inhibition of the methyltransferase activity of PRMT5 occurred in a rapid, mRNA independent manner.
These results implicate PRMT5 as an oncogenic driver that post-translationally regulates oncogenic proteins such as c-myc and cyclin D1 in the context of mantle cell lymphoma.
Since we had adequately established that inhibition of the methyltransferase activity of
PRMT5 caused a decrease in the expression of oncogenic genes and proteins next we wanted to examine the expression of tumor suppressor genes and proteins. We were able to demonstrate the de-repression of ST7, C/EBPβ, and PDCD4, tumor suppressors relevant to PRMT5, cyclin D1, and c-myc respectively, with the use of CMP5 in various
MCL cell lines. Specifically, inhibition of PRMT5 methyltransferase activity increased the protein and mRNA expression of these tumor suppressors and caused a decrease in the recruitment of PRMT5 to the promoters of ST7 and C/EBPβ. Finally, we were able to show that CMP5 worked synergistically with 5-aza and TSA, drugs that target the epigenetic modifiers DNMTs and HDACs respectively. Notably, sub-lethal doses of combination 5- aza, TSA and CMP5 resulted in more apoptosis and a decrease in cyclin D1 and symmetric dimethyl-H4R3 protein expression.
The t(11; 14)(q13;q32) translocation is a distinct feature of MCL that results in the overexpression of cyclin D1, highlighting the importance of cell cycle disruption in this disease. This chapter demonstrated that the epigenetic modifier PRMT5 is over- expressed in MCL and directly associates with cyclin D1. Attenuation of PRMT5
104 methyltransferase activity decreased the expression of malignant proteins and increased the expression of tumor suppressors. Further investigation is needed to completely characterize the relationship between PRMT5 and cyclin D1 and to ascertain the mechanism of PRMT5-controlled cell cycle in MCL, however evidence in this study points to a post-translational mechanism of control. Particularly, it is unclear which oncogenic protein is upregulated first in MCL. We theorize that the translocation results in the upregulation of cyclin D1 which then upregulates PRMT5, which assists in the stabilizing and efficient shuttling of cyclin D1 protein, perpetuating the malignancy.
In this chapter we also described the use of PRMT5 inhibitors in in vitro models of MCL.
These inhibitors allowed us to develop a cursory view of the role of PRMT5 in MCL.
Attempts to use these inhibitors in an in vivo model of MCL, generated by engrafting
SCID mice with MCL cell lines, failed because the study had to be stopped due to toxicity. Further work will have to be performed to develop more potent inhibitors with lower IC50 values and to develop a better delivery system for future in vivo use. In the future these inhibitors could be used to study the mechanism of PRMT5 dysregulation in various malignancies. Most importantly, the synergy experiments outlined in this chapter indicate that PRMT5 inhibitors could potentially be used in the clinic as a part of a combination chemotherapeutic strategy.
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Chapter 3: PRMT5 transgenic mice develop lymphomas
Introduction
Non-Hodgkin’s lymphoma (NHL) is a complex set of cancers that affect the lymphocytes in the blood, the majority of which occur in B-cells. The five year relative survival for NHL as a whole has improved to 69%, due largely in part to increased knowledge about the disease translated into the formulation of new clinical drugs. Despite this improvement in survival, subsets of NHL are aggressive and refractory to current treatment options and thus would benefit from new cancer therapeutics. In order to design and create more potent therapeutics it is necessary to use animal models that mimic the disease of interest. Animal models of disease allow researchers to closely follow disease progression, potentially discover new targets to use for drug development and test drugs of interest in the disease model before they are released to the general public.
Animal models typically begin in the mouse and progress to more complex animals as trials prove successful or more mechanistic information is needed. Inducible mouse
(murine) models of lymphoma tend to engraft human lymphoma cell lines or primary cells into the severe combined immunodeficiency (SCID) mouse, which lacks functional B and T lymphocytes, and therefore does not allow for an accurate assessment of the immune response as it relates to the tumor microenvironment. Immunocompetent models use murine cell lines for engraftment which may not completely mimic the 106 genetic signature and symptoms of disease as the human counterpart. Spontaneous murine models of lymphomagenesis manipulate the genome of mice to either knock-in known oncogenes or knock-out known tumor suppressors and study natural disease progression. While they have the disadvantage of variable time to onset of disease and penetrance of tumor formation, spontaneous models of disease allow researchers to study the contribution of a particular gene or pathway to the development of the malignancy in question.
Several studies have implicated the epigenetic regulator PRMT5 as being a major driver of lymphomagenesis [155,175,177,194,199,237]. Notably, PRMT5 is overexpressed in primary patient lymphoma cells relative to resting and activated lymphocytes.
Additionally, inhibition of PRMT5 prevents Epstein-Barr Virus mediated transformation of
B-cells, suggesting that PRMT5 is a crucial element needed for lymphocyte transformation. We hypothesized that over-expression of PRMT5 represents a transforming event that promotes lymphoma. In order to ascertain the role of PRMT5 in the development of lymphomagenesis we generated a PRMT5 transgenic mouse that over-expressed human PRMT5 (hPRMT5) in B-lymphocytes via the Eµ promoter. This chapter describes the characterization of a PRMT5 transgenic mouse model and offers insights into the mechanism of PRMT5-driven lymphomagenesis.
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Materials and Methods
Molecular cloning
PCR primers to amplify full-length hPRMT5 were designed with loxp sites flanking the sequence followed by flanking EcoRI sites. An initial PCR reaction was performed with the primers using cDNA from 293Ts and Taq polymerase (Thermo Fisher Scientific, Grand
Island, NY). PCR conditions were as follows: Initialization at 95°C for 4 minutes, and 30 cycles of denaturation at 95°C for 1 minute, annealing at 45°C for 30 seconds, elongation at 68°C for 3 minutes followed by a final elongation step at 68°C for 3 minutes. The PCR product was separated on a 1% agarose gel and gel purified using the QIAquick® gel extraction kit (Qiagen, Valencia, CA). The purified product was sent for Sanger sequencing at the Ohio State University sequencing core to verify the correct product was obtained. Once verified the purified PCR product was ligated to the pBH vector via the BglII site using TOPO TA kit for ligation (Thermo Fisher Scientific Grand Island, NY).
The ligated vector was then cloned into TOP10 competent E. coli using the TOPO® TA cloning ® kit from (Thermo Fisher Scientific, Grand Island, NY). The cloning mixture was streaked onto agar plates that contained 100 µg/mL ampicillin sodium salt (Fisher
Scientific, Pittsburgh, PA) and allowed to incubate overnight at 37°C. The following day several ampicillin-resistant colonies were picked to be incubated in LB broth with 100
µg/mL ampicillin sodium salt (Fisher Scientific, Pittsburgh, PA) at 37°C in a shaking incubator overnight. Purified plasmid was isolated from the broth using the QIAprep®
Spin Miniprep kit (Qiagen, Valencia, CA). The purified plasmid was separated on a 1%
108 agarose gel and gel purified as above using the QIAquick® gel extraction kit. Sanger sequencing was used to confirm the presence of the vector and the correct orientation of the hPRMT5. The primers used were: PRMT5 loxp BglII primer1: 5’ – GCG CAG ATC
TAT AAC TTC GTA TAG CAT ACA TTA TAC GAA GTT ATA AGA TGG CGG CGA TGG CGGT
– 3’ and PRMT5 loxp BglII primer2: 5’- GCG GAG ATC TAT AAC TTC GTA TAA TGT ATG
CTA TAC GAA GTT ATC TAG AGG CCA ATG GTA TAT GAGCG-3’ After confirmation of the presence of the vector the product was transformed into TOPO10 competent E. coli and allowed to incubate overnight on agar plate with 100 µg/mL ampicillin sodium salt
(Fisher Scientific, Pittsburgh, PA). Ampicillin-resistant colonies were picked the next day, placed in broth with 100 µg/mL ampicillin sodium salt (Fisher Scientific, Pittsburgh, PA) and grown overnight at 37°C in a shaking incubator. The next day the plasmid was purified from the bacteria grown in the broth with the QIAprep® Spin Miniprep kit
(Qiagen, Valencia, CA). The purified plasmid was then digested with BamH1 (Thermo
Fisher Scientific, Grand Island, NY) to release the pure vector of interest. The purified plasmid was separated on a 1% agarose gel and gel purified as above using the
QIAquick® gel extraction kit. Sanger sequencing was used to confirm the presence of the vector hPRMT5. After confirmation the OSUCCC Transgenic Core Facility performed pronuclear injections of the EcoRI fragment released from the pBH-hPRMT5 vector into
0.5 day old fertilized eggs from FVB/N mice (Taconic, Hudson, NY).
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Housing of mice
All animal experiments were carried out under protocols approved by the Ohio State
University, University Institutional Laboratory Animal Care and Use Committee (ULAR)
(Animal Welfare Assurance #A3261-01). Mice were housed in a ULAR approved vivarium and supplied with sterile food and water ad libitum.
Murine B- and T-cell lymphoma transfection
EL4 and WEH1 cells were transfected with 2 µg of either control, empty vector or pBH-
PRMT5 plasmid using an Amaxa Nucleofector® Kit and apparatus (Lonza, Allendale, NJ).
The Nucleofector® Kit L was used for both the EL4 and WEH1 cell lines and the transfection proceeded by the instructions included in the kit.
Inserting a single copy of PRMT5 into wild type FVB/N mouse
Tail DNA isolated from a wild type FVB/N mouse was combined, at a concentration of 10
µg, with 4.6 pg of hPRMT5-loxp primers in a PCR reaction with Taq polymerase. PCR conditions were as follows: Initialization at 95°C for 4 minutes, and 35 cycles of denaturation at 95°C for 1 minute, annealing at 50°C for 30 seconds, elongation at 72°C for 3 minutes followed by a final elongation step at 72°C for 10 minutes. The product from the PCR reaction was used as a positive control for the original genotyping of the founders.
Genotyping
DNA was isolated from tail snips via the REDExtract-N-Amp Tissue PCR kit (Sigma, St.
Louis, MO) per the manufacturer’s directions. A positive PRMT5 genotype was confirmed
110 using PCR performed using the reagent in the REDExtract kit and primers for the pBH vector. pBH primers are as follows: sense: AAAACCACTTCTTCAAACCACAGC and antisense: CTGCTGGAGAAGAAGGGACATC.
Protein isolation and quantification
Cell pellets were resuspended in RIPA buffer made with 10mM pH 7.4 Tris (Fisher
Scientific, Pittsburgh, PA) 150 mM NaCl (Fisher Scientific, Pittsburgh, PA), 1% Triton X-
100 (Fisher Scientific, Pittsburgh, PA), 1% deoxycholic acid (Acros Organics [Thermo
Fisher], Pittsburgh, PA ), 0.1% SDS (Gibco [Thermo Fisher Scientific], Grand Island, NY) and 5 mM EDTA (Fisher Scientific, Pittsburgh, PA), with the protease inhibitors Sigma and phenylmethanesufonyl fluoride (PMSF) and the phosphatase inhibitor cocktails 2 and 3
(Sigma Aldrich, St. Louis, MO) added at a 1:100 dilution. Pellets were either immediately frozen at -80°C or allowed to lyse on ice for 10 minutes with vortexing before and after the incubation. After either thawing the pellets on ice or lysing on ice the samples were centrifuged at maximum speed (16,000 x g) for 10 minutes in a refrigerated 5415
Eppendorf microcentrifuge (Eppendorf, Hauppauge, NY). The cell pellets were discarded and the protein concentration of the supernatant was determined via the BCA Protein
Assay Kit (Thermo Fisher, Pittsburgh, PA) per the manufacturer’s instructions. In brief, aliquots of samples were diluted 1:10 in lysis buffer with no protease or phosphatase inhibitors and 10 µL of each were added in triplicate along with the standards to a Falcon
96-well flat bottom plate (Corning, Corning, NY). The Reagent A was mixed in a 50:1 ratio with reagent B and 200 µL of the mix was added to each well. The plate was incubated at
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37°C in a cell culture incubator for 30 minutes. After 30 minutes the plate was allowed to cool on the benchtop for 10 minutes. The absorbance of the plate was read with a
Labsystems Multiskan MCC/340 (Thermo, Vantaa, Finland) absorbance plate reader.
Western Blot
20 µg of protein was combined with an equal volume of 2x laemmli sample buffer made by adding 5% β-mercaptoethanol (Fisher Scientific, Pittsburgh, PA) to a stock 2x laemmli sample buffer (Biorad, Hercules, CA). The samples were then boiled for 5 minutes at
99°C. The amount of 20 µg of protein plus 2X sample buffer was loaded into either a pre-cast TGX any kD gel (Bio-Rad, Hercules, CA) or a Tris-HCl gel made with Tris-HCL and
SDS buffer (National Diagnostics, Atlanta, GA) and 40% acrylamide (BioRad, Hercules,
CA) to the desired percentage. Precision Plus Protein ladder (Bio-Rad, Hercules, CA) was loaded in the first lane of each gel to use as a reference guide for the protein weights.
The gel was allowed to run, at 200 V for a pre-cast gel or 120 V for an in-house made gel, in 1X running buffer (National Diagnostics, Atlanta, GA) until the proteins had sufficiently resolved in the gel. Separated proteins from the gel were transferred onto a
PVDF membrane pre-wet with methanol (Fisher Scientific, Pittsburgh, PA) and soaked in transfer buffer. For the precast gels the transfer buffer was made from the Transblot
Turbo transfer stock buffer (BioRad, Hercules, CA) with 200 proof ethanol (Fisher
Scientific, Pittsburgh, PA). For in-house gels, transfer buffer was made with 38.6 mM glycine, 48.9 mM Tris base, 3.7% SDS and 20% methanol (Fisher Scientific, Pittsburgh,
PA). The membrane and blotting pads were soaked in transfer buffer for 15 minutes at
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4°C prior to the transfer. Proteins were transferred from the gel to the membrane via the
Turboblot Plus for pre-cast gels (Bio-Rad, Hercules, CA) or TransBlot SD cell for in-house made gels (Bio-Rad, Hercules, CA). Blots were either allowed to dry for blocking or blocked with 5% milk in 1X TBST (TBS [National Diagnostics, Atlanta, GA] + 0.1% Tween
20 [Sigma, St. Louis, MO]). Blocked blots were then incubated in primary antibody for either 2 hours at room temperature or overnight at 4°C. Primary antibody was diluted
1:1500 in 5% milk made with 1X TBST. After incubation with primary antibody, blots were rinsed four times with 1X TBST for 10 minutes each time. Blots were then incubated with secondary antibody for 1 hour at room temperature. Secondary antibody was diluted
1:4000 in 5% milk made with 1X TBST. After incubation with secondary antibody, blots were rinsed four times with 1X TBST for 10 minutes each time. Blots were developed utilizing either SuperSignal™ West Pico chemiluminescent substrate (Thermo Fisher,
Pittsburgh, PA) or WesternBright™ ECL (Advansta, Menlo Park, California). If a more sensitive detection method was required SuperSignal™ West Femto chemiluminescent substrate (Thermo Fisher, Pittsburgh, PA) or WesternBright™ Sirius ECL (Advansta, Menlo
Park, California) was used. Bands were visualized on Hyblot CL autoradiography film
(Denville, Holliston, MA) developed by a Kodak M6B (TI-BA Enterprises, Inc, Rochester,
NY).
Real-time (RT) quantitative polymerase chain reaction (PCR) assay
Total RNA was prepared from cells using TRIzol reagent (Thermo Fisher Scientific, Grand
Island, NY) according to the manufacturer's instructions. cDNA was synthesized using the
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Taqman MicroRNA Reverse Transcriptase Kit (Thermo Fisher Scientific, Grand Island, NY) per the kit’s directions. Real-time PCR was performed in triplicate for each sample using
TaqMan 2X Universal PCR Master Mix (Thermo Fisher Scientific, Grand Island, NY) using the manufacturer’s directions. MicroRNA expression levels were quantified using the
Applied Biosystems 7900HT Fast Sequence Detection System (Carlsbad, CA). Taqman
PRMT5 primers (product number: 4351370; assay ID: Hs01047356_m1) were used for real-time PCR and GAPDH Taqman primers (product number: 4331182; assay ID:
Mm99999915_g1) were used as a normalization control (Thermo Fisher Scientific, Grand
Island, NY). Expression levels were verified by three independent experiments.
Cell Surface flow cytometry
Single cell suspensions were prepared from spleen, liver and blood where applicable.
Residual red blood cells were lysed with a lysis buffer consisting of 150 mM ammonium chloride, 10 mM potassium bicarbonate and 0.09% EDTA. About half a million cells were resuspended in 1X PBS, pH 7.4 ((Gibco, Grand Island, NY) per condition. The cells were stained with the appropriate antibodies for 30 minutes on ice in the dark. All antibodies were obtained from BD Biosciences (San Jose, CA). After 30 minutes, cells were washed with 1X PBS, pH 7.4 and resuspended in fresh PBS. Cells were analyzed via a FC500
Beckman Coulter flow cytometer (Beckman Coulter, Pasadena, CA).
B, NK and T-cell sorting
A single cell suspension of splenocytes isolated from non-transgenic and transgenic mice was used to sort for B, NK and T-cells. Cells were gated and sorted on B220 FITC for B-
114 cells, NK1.1 APC for NK cells and CD3 PE for T-cells. The results from three mice for each group were pooled for the final results.
Histology
All tissues were placed in 1% formalin immediately after harvesting. Tissues were paraffin-embedded by The Ohio State University CCC Comparative Pathology and
Mouse Phenotyping core facility. Sections were also cut from the paraffin blocks and stained with H&E by the core facility.
Development of Cell Line from Transgenic Mouse Tumor
At approximately 5 months of age, mouse number 813 began to present with baldness on the face and hyperactivity. One month after the initial symptoms, 813 presented with difficulty breathing and a hunched posture and the veterinarians determined that she had met early removal criteria. Upon euthanization, it was discovered the mouse had a large tumor filling the chest cavity. The tumor tissue was passed through a cell strainer to obtain a single cell suspension and remove unwanted fibrous tissue. Cells were then placed in U-bottom 96-well plates at a concentration of 1 x 106cells/well and cultured in the presence of RMPI (Gibco, Grand Island, NY), 20% FBS, 1X anti-anti (Gibco, Grand
Island, NY) and 1X penicillin/streptomycin (Gibco, Grand Island, NY). Cells were examined daily for viability and potential growth. Cells were split 1:1 once a week as they began to utilize the nutrients in the cell culture media. Within 1 month a viable suspension cell culture was obtained.
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813 cell culture
The semi-adherent 813 cell line was maintained in RMPI without glutamine (Gibco,
Grand Island, NY) supplemented with 10% heat inactivated fetal bovine serum (FBS)
(Atlanta Biologicals, Norcross, GA or Sigma, St. Louis, MO) 1X glutamax (Gibco, Grand
Island, NY) and 1X penicillin/streptomycin (Gibco, Grand Island, NY). Cells were kept at
37°C in a Thermo 3110 incubator (Fisher Scientific, Pittsburgh, PA) with 5% CO2 concentration. Cells were subcultured every 2-3 days at a concentration of 2 x 105 cells/mL. TrypLE™ Express (Thermo Fisher Scientific, Grand Island, NY) was used to dislodge adherent cells from the flask. Cell counting was performed utilizing a hemocytometer and trypan blue staining to visualize dead cells.
Human PRMT5 PCR analysis
RNA was isolated according to the trizol (Thermo Fisher Scientific, Grand Island, NY) protocol. cDNA was prepared using the MMLV Reverse Transcription Kit (Thermo Fisher
Scientific, Grand Island, NY) and following the manufacturer's recommendations. PCR primers were designed to recognize human PRMT5 and not mouse PRMT5. Primer sequences are: forward GCTGCCCCTTAATCAGGAAG and reverse
ACGTACCGTTATGGGCTGC and were ordered from Operon with a salt-free purification. A
PCR master mix was made utilizing Platinum® Taq DNA polymerase (Thermo Fisher
Scientific, Grand Island, NY) and following the instructions supplied with the Taq. PCR cycling conditions are as follows: hold for 94°C, denature at 94°C for 30 seconds, anneal at 59°C for 45 seconds and extend at 72°C for 1.5 minutes. The denature, anneal and
116 extend steps were repeated for 30 cycles with a final extension step at 72°C for 10 minutes. DNA was separated in a 1.2% agarose gel made with 0.5 mg/mL ethidium bromide (Thermo Fisher Scientific, Grand Island, NY) and run for about 2 hours at 100V.
Loxp and pBH PCR and Sanger sequencing
RNA was isolated by Trizol (Thermo Fisher Scientific, Grand Island, NY) and cDNA was made using the MMLV Reverse Transcription Kit (Thermo Fisher Scientific, Grand Island,
NY) following the manufacturer's recommendations. PCR was performed using
Platinum® Taq DNA polymerase (Thermo Fisher Scientific, Grand Island, NY) and following the protocol outlined in the product insert. For a 50 µL reaction mixture used
200 µM each of the forward and reverse primers and 1 µL of cDNA. GAPDH primers used were: forward 5’ – TTCCATCCTCCAGAAACCAG – 3’ and reverse 5’ –
CCCTCGAACTAAGGGGAAAG – 3’. Loxp primers used were: forward 5’-
AGCCAGAACCGTCCTCCACCTAA – 3’ and reverse 5’ – GGAAGCGTTCACCAGGGGTCC – 3’.
The following parameters were used for the loxp primers: 95°C for 15 minutes, then 34 cycles of 95°C for 30 seconds, 55.3°C for 30 seconds and 72°C for 60 seconds, followed by a final 10 minute annealing step at 72°C. The following parameters were used for the pBH vector primers: 94°C for 3 minutes, then 33 cycles of 95°C for 45 seconds, 60°C for
30 seconds and 72°C for 45 seconds, followed by a final 10 minute annealing step at
72°C. PCR products were mixed 1:10 with 10X BlueJuice™ Gel Loading Buffer (Thermo
Fisher Scientific) and run on a 1.2% DNA gel at 100 volts. The bands of interest were cut out of the gel and digested using the QIAquick Gel Extraction Kit (Qiagen, Valencia, CA).
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Catalog Antibody Clone Fluorochrome Number B220 RA3-6B2 553092 APC CD3 17A2 555274 FITC CD3 17A2 555275 PE CD4 RM4-5 553051 APC CD4 RM4-5 553048 PE CD5 53-7.3 553022 PE CD8a 53-6.7 553030 FITC CD19 1D3 553786 PE CD24 M1/69 553262 PE CD25 PC61 557192 APC CD43 S7 553271 FITC CD45 30-F11 559864 APC CD69 H1.2F3 553237 PE CD117 2B8 553356 APC CD122 5H4 554452 FITC IgM 11/41 553437 FITC NK1.1 PK136 557391 PE Table 6. Antibodies used for flow cytometry.
To prepare the sample for Sanger sequencing, about 15 ng of DNA was diluted in 12 µL
of RNA, DNAse free H2O and 6.4 pmol of the forward primer. The same procedure was repeated for the reverse primer. The samples were then submitted to the OSU Core for
Sanger sequencing.
Flow cytometry
About 1 million cells per condition were used for the surface staining of cytokines of interest. Cells were pelleted by centrifugation for 5 minutes at 1500 rpm and resuspended in 500 µL of 1X PBS, pH 7.4 along with 0.5 µL of the antibody of interest.
The stained cells were incubated in the dark for 30 minutes on ice. After the incubation
118 the cells were washed with 1 mL of 1X PBS, pH 7.4 and pelleted by centrifugation. The cells were then resuspended in 500 µL of 1X PBS, pH 7.4 and run immediately on the flow machine or fixed in 200 µL 1% formalin and run at a later time. Antibody information is located in Table 6.
Intracellular flow cytometry
About 1 million cells per condition were used for the initial surface staining of cytokines of interest. Cells were pelleted by centrifugation for 5 minutes at 1500 rpm and resuspended in 500 µL of 1X PBS, pH 7.4 along with 0.5 µL of the antibody of interest.
Conditions were included for isotype controls for all of the antibodies used. The stained cells were incubated in the dark for 15 minutes on ice. After the incubation the cells were washed with 1 mL of 1X PBS, pH 7.4 and pelleted by centrifugation. The cells were then resuspended in 250 µL of BD Cytofix/Cytoperm buffer (Fisher Scientific, Pittsburgh, PA) to fix and permeabilize the cells. The cells were vortexed and incubated covered on ice for
20 minutes. The cells were then washed twice with 1 mL 1X Perm/Wash buffer (Fisher
Scientific, Pittsburgh, PA) to keep the cell permeabilized during the washing steps and pelleted by centrifugation between washes. The cells were then resuspended in 100 µL perm/wash buffer and stained for intracellular cytokines with 0.5 µL of the desired antibody per tube. The cells were incubated covered on ice for 30 minutes. After the 30 minute incubation the cells were washed once with 1 mL 1X Perm/Wash buffer and pelleted by centrifugation. The cells were then resuspended in 250 µL 1% formalin and used to perform flow cytometry. Antibody information is located in Table 6.
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T-cell Receptor (TCR) Analysis
Total RNA was isolated from early and late passage 813 cells by TRIzol reagent (Thermo
Fisher Scientific, Grand Island, NY) according to the manufacturer's instructions. cDNA was prepared with the MMLV Reverse Transcription Kit (Thermo Fisher Scientific, Grand
Island, NY) following the manufacturer's recommendations. Vβ TCR expression was analyzed via PCR using the TCRExpressTM kit (BioMed Immunotech, Tampa, FL). PCR products were separated with a high resolution 4% gel made from components supplied in the kit. Ethidium bromide (Thermo Fisher Scientific, Grand Island, NY) was added to the gel at a concentration of 0.5 mg/mL. The gel was run for about 2 hours at 100V. The bands in the gel were visualized with a Fluorchem E imaging system (ProteinSimple, San
Jose, CA).
Cytogenetics
Exponentially growing cells were fixed using standard laboratory procedures. The cell suspension was dropped onto precleaned, warm, wet, slides. The slides were aged at
90oC for 1 hour, banded with trypsin and stained with Wright stain. Banded metaphases were analyzed using an Axioskop 40 (Zeiss, Thornwood, NY). For each cell line at least 20 metaphases were karyotyped using an Applied Imaging Karyotyping System.
T-cell activation
Functional grade mouse Anti-CD3 antibody (eBioscience, San Diego, CA) was resuspended at a concentration of 5 µg/mL in sterile PBS. Tissue culture plates were coated with the anti-CD3 antibody (1 mL in 6-well plates) and incubated overnight at
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4°C. The following day the cells of interest were resuspended at a concentration of 2 x
106 cells/mL, in fresh media, along with 1:1000 CD28. The excess plate bound anti-CD3 was removed and 2 mL of cells were added per well in a 6-well plate. The cells were incubated for 24 hours at 37°C. T-cell activation was measured by an increase in surface
CD25 and CD69 analyzed by flow cytometry.
MTS Assay
813 cells were split the day before to 2 x 105 cells/mL. The day of treatment 100 µL of cells were plated into a 96-well round bottom plate. 100 µL of drug diluted in media was added to the cells for a final volume of 200 µL/well. Drug concentrations were initially doubled to account for the final 1:1 dilution within the well. All treatments were performed in quadruplicate within the plate and 4 wells per plate were filled with 200 µL of media only to serve as blanks. Cells were incubated for the desired time length. Cell proliferation was measured using the CellTiter 96® AQueous Non-Radioactive Cell
Proliferation Assay (Promega, Madison, WI). The MTS and PMS solution was mixed in a
20:1 ratio and 25 µL of this mixture was added to each well. The plate was allowed to incubate for 2-4 hours at 37°C. After the desired incubation time, the absorbance was read at 490 nm using a Labsystems Multiskan MCC/340 (Thermo, Vantaa, Finland) absorbance plate reader. The blank absorbance values were subtracted from the absorbance values of the samples.
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Annexin V/PI flow cytometry
An equal amount of cells for each condition were pelleted by centrifugation for 5 minutes at 1500 rpm. After centrifugation the supernatant was removed and resuspended in 100 µL of a master mix containing 100 µL 1X Annexin Binding Buffer (BD
Biosciences, San Jose, CA) with 5 µL FITC Annexin V (BD Biosciences, San Jose, CA) and 5
µL propidium iodide (BD Biosciences, San Jose, CA). The cells were allowed to stain for 15 minutes. After 15 minutes, an additional 400 µL of 1X Annexin Binding Buffer was added to the cells. Cell viability was then checked via a FC500 Beckman Coulter flow cytometer
(Beckman Coulter, Pasadena, CA).
813 engraftment
The appropriate number of cells were centrifuged and resuspended in 200 µL PBS pH
7.4. Cells were injected via tail vein using 31 gauge insulin needles (BD, Franklin Lakes,
New Jersey) into either CB.17 SCID or FVB/N mice (Taconic, Hudson, NY).
Crispr-Cas9 PRMT5 knockout
A dual vector lentiviral system was designed to consist of a constitutive expression vector for the Cas9 endonuclease, a doxycycline (dox)-inducible single guide RNA (sgRNA) cassette and an ubiquitin promoter driven tetracycline repressor linked via the T2A peptide to the GFP promoter. This system was a gift from Dr. JM Herold, Department of
Medical Biology, University of Melbourne, Parkville, Australia (Cell Reports, 2015, PMID
25732831). sgRNA were designed by utilizing the MIT CRISPR design software
(http://crispr.mit.edu). For the transfection lenti-X 293Ts (Clontech, Mountain View, CA)
122 were plated 5 x 106 cells per 10 cm tissue culture plate in antibiotic-free DMEM media one day prior to transfection. On the day of the transfection the media on the 293T cells was changed to fresh complete RPMI + 10% FBS media with antibiotics and glutamax 3 hours prior to the transfection. 293T cells were transfected with the lentiviral Cas9-EGFP- blasticidin resistant plasmid and the lentiviral packaging plasmids pCMV and VSV-G, utilizing Opti-Mem® media and Lipofectamine® 2000 (Thermo Fisher Scientific, Grand
Island, NY) transfection reagent and following the procedure provided with the reagent.
The transfection was allowed to continue for 72 hours after which time the viral particles were harvested by filtering the 293T supernatant that contained the virus with a 0.45 micron filter (EMD Millipore, Billerica, MA.). Tg813s split the day before to 2 x 105cells/mL in fresh complete RPMI media were concentrated to 5 x 106cells/mL in fresh complete
RPMI media, plated in 6-well plates with 1 mL of cells per 1 mL of virus along with 10
µg/mL polybrene (Sigma Aldrich, St. Louis, MO) and allowed to incubate for 30 minutes at 37°C. After the 30 minute incubation the plates were centrifuged for 2.5 hours at 2500 rpm (spinoculation). After the centrifugation the cells were incubated for 14 hours at
37°C after which the media was aspirated and the cells were placed in fresh complete
RPMI and allowed to incubate for up to 72 hours after the media change. GFP expression was checked by flow cytometry every 24 hours during this time until GFP expression was greater than 90%, after which time the cells were placed under blasticidin selection and single cell cloned by serial dilution in 96-well plates. Cas9 expression was confirmed by
123 immunoblotting using anti-CRISPR-Cas9 antibody. PRMT5 knockdown was induced by treating cells with 1 µg/mL fresh doxycycline dissolved in sterile water.
Statistical Analysis
Results were expressed as the means ± S.D. unless otherwise specified. F-tests for analysis of variance and paired t-tests were conducted in Microsoft Excel 2010 and used to generate p-values for comparisons between two groups and when multiple samples within different groups were used, respectively. Kaplan Meier analysis for engraftment models was performed in JMP Pro 12.
Results
Establishment of the hPRMT5 model
We hypothesized that PRMT5 is
required to initiate and maintain
B-cell lymphomagenesis thus we
sought to create a transgenic
Figure 29. Schematic of Prmt5 vector. mouse model that over- Human PRMT5 cDNA, flanked by loxp sites, was cloned into a pBABE-Hygro vector under control of the Eµ promoter. expressed human PRMT5 in
murine B-cells. To target lymphoid cells for human PRMT5 (hPRMT5), the floxed gene was introduced into the pBABE-hygro (pBH) vector containing the IgH Enhancer/Promoter (Eµ), and polyadenylated sequence (Figure 29). To confirm the ability of the fragment to drive overexpression of PRMT5 in lymphocytes, the EcoR1 digested fragment was transfected
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Figure 30. The pBH vector containing hPRMT5 can be efficiently transfected into EL-4 and WEH1 mouse T and B-cell lymphoma cell lines respectively.
A) GFP expression in EL-4 and WEH1 cells after transfection with GFP-tagged scramble plasmid was over 30%. B) Western blot analysis of protein expression of PRMT5 in EL-4 and WEH1 transfected cells show an increase in PRMT5 protein in transfected cells relative to untransfected or vector transfected cells. No vector = 0, empty vector = pBH and vector with hPRMT5 = PRMT5 C) Prmt5 mRNA expression in EL-4 and WEH1 cells transfected with hPRMT5 plasmid is increased as analyzed by quantitative real-time PCR analysis. *p<0.05, **p<0.01 compared to empty vector control.
into the mouse B and T cell lymphoma cell lines, WEH1 and EL-4 respectively. Scramble- tagged GFP was transfected in parallel with the pBH-PRMT5 plasmid and transfection efficiency was greater than 30% in both cell lines as measured by flow cytometry
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Figure 31. PCR analysis of the pBH vector in the original group of pups obtained after pronuclear injections of the pBH vector containing the human PRMT5 transgene reveals 5 potential founder mice.
DNA was isolated from tail snips from each of the pups represented by the numbers across the top. The far left panel on the second row is over-exposed to visual the positive control (PC) which is DNA from a wild type FVB/N mouse with 1 copy of Prmt5 inserted. The negative control (NC) is DNA from a wild type FVB/N mouse. The PCR analysis yielded 5 founders (15, 20, 24, 30, and 31) that displayed robust expression of the transgene.
(Figure 30A). More importantly, the hPRMT5 plasmid significantly increased the protein and mRNA expression of PRMT5 in both WEH1 and EL-4 cell lines (Figure 30B and
Figure 30C). After confirming that the plasmid could increase PRMT5 transcript and protein in in vitro model, we utilized the services of the Transgenic Facility at The Ohio
State University Comprehensive Cancer Center to perform pronuclear injections of the hPRMT5 containing vector into 0.5 day old FVB/N embryos. From this injection we obtained 36 potential founder mice of which 5 mice, numbered 15, 20, 24, 30 and 31, contained the transgene, as analyzed by PCR using primers specific for the pBH vector
(Figure 31). Mouse number 33 also contained the vector however copy number analysis
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Figure 32. The 5 candidate founder mice express hPRMT5.
A) by PCR and protein B) by western blot. C) Real time PCR analysis of human PRMT5 expression in splenocytes (15, 24 and 31) or bone marrow (30) from F1 generation of PRMT5 transgenic mice. Experiment was from 1 set of samples from each founder analyzed in quadruplicate. nTg = non-Transgenic; Tg = Transgenic * p-value < 0.001 compared to non-transgenic mice.
revealed that it contained significantly fewer copies than the other 5 founder mice thus it was not used for subsequent breeding (data not shown). Repeat DNA PCR analysis in these mice revealed a high expression of the pBABE-hygro vector containing hPRMT5 as compared to non-transgenic mice (Figure 32A) and protein from splenocytes isolated
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Figure 33. The baseline immunophenotype does not differ between non-transgenic and transgenic mice.
There is no difference in the CD19+ population in A) unpurified splenocytes or B) bone marrow between non-transgenic and transgenic mice. Single-cell suspensions of splenocytes and bone marrow were isolated from non-transgenic or transgenic F1 mice from founder 30. Two mice were used for each non-transgenic or transgenic condition. Representative flow is shown from 1 set.
from Founder 30 at the time of its death also confirmed the overexpression of PRMT5 128
Figure 34. Prmt5 mRNA is over-expressed in the lymphocytes of transgenic mice.
Splenocytes were isolated from 3 transgenic or non-transgenic mice and sorted into B, NK and T-cell groups based on their immunophenotype as evaluated by FACS flow cytometry. RNA from the sorted B, NK and T- cells was isolated and converted to cDNA. Prmt5 mRNA expression was determined by quantitative real time PCR and pooled for the 3 samples. Prmt5 mRNA is overexpressed in the B, NK and T-cell compartments of hPRMT5 transgenic mice as compared to non-transgenic age matched controls.*p-value <0.05; **p-value <0.01 N = 3 for NK and T cells and 2 for B cell transgenic.
protein in the tissues of the mouse as compared to a non-transgenic control mouse
(Figure 32B). The 5 founder mice were mated with wild type (WT) FVB/N mice in order
to obtain an F1 generation. Representative F1 pups from each founder that were positive
for the transgene, overexpressed hPRMT5 mRNA in their spleens and bone marrow as
compared to non-transgenic controls, indicating that transgene expression was heritable
(Figure 32C). Founder 15 never developed a measureable phenotype of considerable
penetrance which could be due to a lower copy number expression of hPRMT5 as
compared to the other founders as demonstrated by the small amount of PRMT5 mRNA
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Figure 35. A small subset of transgenic mice develop lymphoma-like symptoms.
Representative pathology of hPRMT5 transgenic mice. Founder 30 presented with hepatomegaly, nephromegaly and splenomegaly relative to an age matched non-transgenic mouse.
in the splenocytes of the PRMT5 positive F1 mouse from this founder. Additionally, we suspect that the original founder presented with mosaicism due to the low heritability of the transgene in this line.
Baseline phenotypic characteristics of hPRMT5 transgenic mice
There were no physical phenotypic differences between transgenic and non-transgenic mice. The PRMT5 transgenic mice also displayed no significant differences in surface
CD19 expression in either their spleens or bone marrow as compared to non-transgenic animals of comparable age as determined by flow cytometry (Figure 33A and Figure
33B). In order to verify the presence of the hPRMT5 transgene in the lymphocyte compartment, the mRNA levels of Prmt5 were examined in B, T and NK cells sorted from
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Figure 36. Founder 30 displays lymphocyte infiltrate in various organs and tissues as compared to an age matched non-transgenic control.
A) Peripheral blood smears show a moderate increase of lymphocytes, indicated by the arrows, in founder 30 versus a non-transgenic mouse. B) H&E stains of the liver, lung, spleen, and gastrointestinal tract (G.I.) tract of founder 30 versus show an increase in lymphocyte infiltrate in these organs concurrent with ablation of the normal architecture as compared to a non-transgenic mouse.
non-transgenic and transgenic mice splenocytes. As anticipated, there was significantly more Prmt5 mRNA in the B cells of transgenic mice as compared to non-transgenic mice
(Figure 34). Unexpectedly, there was also significantly more Prmt5 mRNA in T and NK cells of transgenic mice as compared to non-transgenic mice. This indicated that although, the vector did successfully drive the expression of hPRMT5 predominately in B- cells, it was promiscuous and allowed for the transgene expression to leak to the other lymphoid compartments.
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Figure 37. Immunophenotype of Founder 30.
Splenocytes were isolated from Founder 30 and an age matched non-transgenic control mouse and stained for the surface antigens and analyzed by flow cytometry.
Characteristics of disease in hPRMT5 transgenic mice
Mice were allowed to age and develop disease naturally and besides lymphoma the
most common morbidity among these mice was obstructive uropathy. Mice were
euthanized when they presented with some combination of hunched posture, rough
coat, tachypnea, weight loss and general inactivity. Upon necropsy, a diagnosis of
lymphoma was suspected with the presence of splenomegaly and/or hepatomegaly,
nephromegaly, and lymphadenopathy. To confirm the existence of lymphoma, each sick
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Figure 38. F1 pup (#253) from founder 24 presented with disseminated disease.
A) F1 pup (#253) from founder 24 presented with disseminated disease in the liver. B) Immunohistochemical analysis of the spleen revealed a high percentage of PRMT5 (red) positive cells relative to the non-transgenic control (#254). Nuclei were stained blue.
transgenic mouse was euthanized along with a non-transgenic mouse of comparable age and sex, if possible, to serve as a comparative control and subjected to immunophenotypic analysis by flow cytometry and/or immunohistochemistry. Blood, bone marrow, and splenocytes were collected from each mouse to be used for analysis and all other organs, including a specimen from the spleen, were collected and kept in formalin. The collected blood was used for blood smears and flow cytometry analysis, while single cell suspensions of bone marrow and splenocytes were used for immunophenotyping by flow cytometry. Several of the mice developed disease including founder 30 who presented with a rough coat and hunched posture at about 21 months of age and was euthanized. Necropsy determined that Founder 30 had splenomegaly, hepatomegaly and nephromegaly as compared to a non-transgenic control mouse
(Figure 35). Founder 30 also presented with lymphocyte infiltrate in the blood
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(Figure 36A), liver, lung, spleen
and gastrointestinal tract (Figure
36B) which had destroyed the
normal architecture of these
tissues. Immunophenotypic
analysis was conducted using
Figure 39. Mouse #253 presented with an immature B-cell CD19, and B220, to assess for B- lymphoma.
Splenocytes were isolated from transgenic mouse #253 and cells, CD3, CD4 and CD8 to assess age-matched non-transgenic mouse #254 and stained with antibodies for the B-cell surface antigens CD19, B220, and IgM. for T-cells and IgM and CD43 to Analysis by flow cytometry revealed the transgenic mouse had a large population of IgM cells that were also CD19 positive indicating these were immature B-cells. assess immature B and
lymphocyte markers respectively.
Flow cytometric analysis of the unpurified splenocytes did not a reveal a clear phenotype of disease in founder 30 (Figure 37). Additionally, instead of the expected accumulation of lymphocytes in the spleen there appeared to be less lymphocytes relative to the non- transgenic mouse. The lack of apparent lymphocyte accumulation in the spleen could be due to the lymphocytes homing to other secondary lymphoid organs. As no other organs were examined it is unclear exactly where the lymphocytes were concentrated.
There was also no indication of abnormal lymphocyte accumulation in the bone marrow
(data not shown).
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A F1 pup #253 from founder 24
also presented with hunched
posture and was euthanized
around 4 months of age along
with the age-matched non-
transgenic mouse #254. The
Figure 40. hPRMT5 transgenic mice over-express PRMT5 mouse displayed visual protein.
Protein was isolated from splenocytes of representative F2 symptoms of lymphoma progeny from each of the founders and examined for PRMT5, p65, cyclin D1, D3 and E and symmetric dimethyl-H4R3 expression. While hPRMT5 transgenic mice do overexpress including enlarged lymph nodes PRMT5 relative to the non-transgenic control there are no other discernible patterns of expression for the proteins analyzed and and infiltrate in the liver and none of the mice express cyclin D1, p53 or Rb. spleen (Figure 38A). Moreover,
immunohistochemical analysis of splenic tissue revealed a high expression of PRMT5 protein in this tissue and an increase in lymphocyte infiltrates. (Figure 38B).
Immunophenotypic analysis of unpurified splenocytes revealed that the transgenic mouse had a population of immature B-cells that were IgM and CD19 positive (Figure
39) thus we concluded that this mouse had developed an immature B-cell lymphoma.
Penetrance of model
There was a wide range of transgene penetrance for the model within the 5 founder lines
(Table 7). The total number of progeny from each founder is indicated by the “total number of mice” column and the total number of progeny positive for the transgene is indicated by the “total # of transgenic mice” column. Founder 15 displayed the lowest
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Table 7. Penetrance of hPRMT5 murine model. heritability of the transgene as only about 6% of the progeny from that founder confirmed to possess the transgene in contrast to almost 50% for founders 24 and 30 as indicated by the heading “% transgenic”. The absolute numbers of transgenic mice that developed lymphoma is denoted by the label “# of transgenic mice with lymphoma” while the percentage of transgenic mice that developed lymphoma is denoted “% transgenic mice with lymphoma”. The age to onset of lymphoma, indicated as “average age of lymphoma onset”, varied between the founders and ranged from 3 to 26 months.
Transgenic mice from founder 24 had the highest incidence of lymphoma among transgenic mice at 34.6%, as opposed to less than 15% for the remaining founders.
Overall the hPRMT5 transgenic mouse is a low-penetrance mouse model, which could reflect the need of PRMT5 to cooperate with other oncogenes to perpetuate a malignant phenotype.
Molecular analysis of transgenic mice
In order to investigate potential differences between transgenic and non-transgenic mice, we examined the protein expression of oncogenic proteins known to be important
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players in tumorigenesis and known to
interact relevant to PRMT5 was analyzed via
western blotting. Splenocytes from F2
generation hPRMT5 transgenic mice or non-
transgenic mice were isolated and used to
make protein lysates. As expected, hPRMT5
transgenic mice expressed more PRMT5
protein relative to non-transgenic FVB/N
Figure 41. Tg813 mouse has lymphoma. mice (Figure 40). FVB/N mice do express
Organs were isolated from a non-transgenic litter mate mouse number 811 and the transgenic endogenous mouse PRMT5 and due to the mouse number 813, formalin-fixed and paraffin- embedded and stained with hematoxylin and high sequence homology between human eosin as described in materials and methods. Mouse #813 had lymphocyte infiltrate in the liver, and mouse PRMT5 protein, most kidneys, lung and tumor tissue.
commercially available PRMT5 antibodies cross-react with both, thus it is not possible to distinguish between human and mouse
PRMT5 protein in these samples. Aside from PRMT5 expression, transgenic mice did not seem to overexpress any of the other proteins analyzed relative to non-transgenic mice.
Additionally, the non-transgenic mouse sample seemed to express more of the PRMT5 histone mark sym-H4R3(2Me) which could be due to the unequal loading of protein as evidenced by the much higher amount of β-actin in that sample as compared to the other samples. Cyclin D1, p53 and Rb, other proteins known to be dysregulated in tumorigenesis, were also analyzed by western blotting but were not expressed in any of
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Figure 42. Tg813 had a rapidly proliferating T-cell lymphoma.
Formalin-fixed paraffin-embedded spleens from the non-transgenic control mouse and the transgenic mouse 813, along with the tumor from 813 were subjected to immunohistochemical analysis. Tissues were stained with the B-cell marker anti-CD20, the T-cell markers anti- CD3, anti- CD4 and anti-CD8, anti-PRMT5, the hematopoietic progenitor cell marker anti-CD34, the pre-lymphoid marker anti-TdT, the T-cell migration and adhesion marker anti-CD99 and the proliferation marker anti-Ki-67. The spleen and tumor tissue of 813 highly expressed PRMT5, CD3 and Ki57, moderately expressed CD4 and little to no CD20 or CD8. While CD34, TdT and CD99 expression differed between the spleen and tumor tissue. Based on these results we concluded the mouse had a highly proliferating CD4+ T-cell lymphoma.
the samples.
Isolation and creation of 813 cell line from a hPRMT5 transgenic tumor
At approximately 5 months of age, mouse number 813, a F3 generation mouse from founder 30, began to present with baldness on the face and hyperactivity. One month after the initial symptoms, 813 developed tachypnea and a hunched posture and was euthanized. Necropsy revealed that the mouse had a large tumorous mass filling in chest
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Figure 43. The 813 cell line expresses PRMT5.
A) PCR analysis using hPRMT5 specific primers on cDNA from 813 primary tumor cells (813T) or splenocytes (813S) compared to splenocytes from 811 non-transgenic (811S) mouse as a negative control or Jeko cells as a positive control. 813 tumor cells and splenocytes express amounts of PRMT5 comparable to malignant Jeko cells while the non-transgenic mouse does not. PRMT5 protein is also overexpressed in the 813 cell line (813C), primary tumor cells and splenocytes as compared to the 811 non-transgenic splenocytes mouse. Early and late passages of the 813 cell line and primary 813 splenocytes also express higher amounts of PRMT5 mRNA relative to splenocytes from the non-transgenic mouse 811. *p-value<0.05; **p-value<0.001
cavity. H&E analysis of the organs of mouse number 813 revealed extensive lymphocyte infiltrate in the liver, kidney, lungs and tumor tissue (Figure 41). Immunohistochemical analyses determined that the spleen and tumor tissue of mouse 813 contained a high expression of PRMT5 and CD3 as compared the non-transgenic litter mate mouse 811, while the tumor itself, and to a lesser extent the spleen, expressed a large amount of
Ki67 indicating this was a highly proliferative lymphoma (Figure 42). A single cell
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suspension was obtained from
the mass, cultured using the
conditions described in the
materials and methods section,
and within 1 month an
Figure 44. The 813 cell line expresses the original pBH vector and loxp sites. immortalized semi-adherent cell
PCR analysis of cDNA from early and late passages of the 813 culture was obtained. RNA cell line confirmed that the cell expresses both the original pBabe-hygro vector used to deliver the transgene as well as the loxp sites flanking the hPRMT5 sequence. The murine MCL cell isolated from the splenocytes of line FC-mMCL (mMCL) was used as a negative control. the 813 mouse and a non-
transgenic mouse 811, the 813 cell line developed from the tumor and the positive
control, the MCL cell line Jeko, was analyzed for the expression of hPRMT5 using PCR
primers designed to be specific for hPRMT5. Splenocytes from mouse 813 and the
subsequent cell line isolated from the tumor of 813 both expressed hPRMT5 while
splenocytes from the non-transgenic control 811 did not (Figure 43A). Whole cell lysate
isolated from 813 splenocytes and tumor cells also expressed more PRMT5 protein
relative to splenocytes from the control mouse 811 (Figure 43B). Quantitative real time
PCR determined that early and late passages of the 813 cell line expressed 2 and 3 fold
more PRMT5 mRNA respectively than splenocytes from the control mouse #811, while
primary splenocytes from the 813 mouse expressed over 27 fold more PRMT5 mRNA
(Figure 43C). To further verify that the 813 cell line expressed the hPRMT5 transgene,
PCR analysis was performed using primers for the hPRMT5-flanking loxp sites and the
140
pBH vector which
contained hPRMT5. A
murine MCL cell line (FC-
mMCL) was used as a
non-expressing control.
We were able to
demonstrate that both
early and late passages
of the 813 cell line
expressed both the Loxp
sites and pBH vector
(Figure 44), which
confirmed that the cell
line expressed the
original transgene
vector. The PCR product
Figure 45. The 813 cell line is a mature T-cell lymphoma cell line. was separated on an
Cells were stained with the indicated antibodies and analyzed for surface expression by flow cytometry to determine their agarose gel, gel purified immunophenotype. The 813 cell line expresses the lymphocyte marker CD45. 813 does not express the B cell markers CD19, IgM or B220, or and subjected to Sanger the NK cell marker NK1.1. It also does not express the myeloid progenitor marker (CD117). The 813 cell line expresses T cell markers sequencing to verify the CD4 and CD3 the latter of which is lost with long-term passage of the cell line. The cell line also expresses the adhesion marker CD24 and CD154.
141
presence of the transgene. This same
experiment was also repeated with
genomic DNA and yielded the same
results.
Immunophenotype of 813 cell line
The 813 cell line was evaluated for the
presence of common surface
lymphocyte markers by flow cytometry
in an effort to determine its Figure 46. 813 cells express intracellular CD3.
Both early and late passages of the 813 cell line express immunophenotype. (Figure 45). The intracellular CD3 as determined by flow cytometry. cell line failed to express B-cell markers such as B220 (CD45R), CD19 and IgM, the NK marker NK 1.1 or the myeloid progenitor cell marker CD-117 (c-kit). The cell line also did not express the hematopoietic stem cell marker Sca-1, or the myeloid progenitor cell marker Gr1 (data not shown). In contrast
813 cells robustly expressed CD4, a marker typically expressed on T-helper cells, and
CD3, the major component of the T-cell receptor. The cells also expressed CD5, which is a B1 and T cell marker, and CD24, an adhesion marker, reflecting its semi-adherent properties. Immature T-cells either lack both CD4 and CD8 or express both, however the
813 cell line expressed only CD4. These data closely mirrored that obtained from the immunohistochemical analysis and it was concluded that the 813 cell line was a mature
T-cell lymphoma cell line. Interestingly, surface expression of CD3 was lost as the cell line
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Figure 47. Early and late passages of the 813 cells express the Vβ17 T-cell receptor subunit.
RNA from early and late passages of the 813 cells was used to examine T-cell clonality by a kit utilizing a nested PCR reaction to analyze Vβ gene families of the T cell receptor. Early and late passages of the cells both displayed high expression of the Vβ 17 family however, the early passage cells were polyclonal while the late passage cells were monoclonal.
was passaged long term for more than 20 passages. In order to further explore what happens to the surface CD3 that is lost as the cell line is passaged, intracellular flow cytometry was employed to examine the intracellular levels of CD3 in both early and late passages of the 813 cell line. We were able to confirm that both early and late passages of the cell line expressed high levels of intracellular CD3 (Figure 46). To ensure that the
813 cell line was of T-cell lineage, T cell clonality was assessed using a nested PCR kit for
Vβ T-cell receptor subunits using RNA from early and late passages of the 813 cell line.
Both early and late passage 813 cells highly express the Vβ 17 family of the TCR
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Figure 48. 813 cells express an abnormal karyotype.
The chromosomal profile of both early and late passages of the 813 cell line revealed abnormalities. A) A representative picture from a clone from the early passage of the cell line demonstrates trisomy in chromosome 6 and 15 and a deletion in chromosome 6. B) A representative picture from a clone isolated from the late passage of the cell line reveals tetraploid and hexaploid chromosomes as well as (15;16) and (4;23) translocations.
(Figure 47). Notably the early passage of the cell line was polyclonal while the later passage of the cell line expressed a monoclonal population of Vβ 17. These experiments
144
indicate that although later
passages of the cell line lose
surface expression of CD3 there is
still a substantial amount of
intracellular CD3 that remains.
Additionally the loss of surface
CD3 expression could be
correlated with the change in
clonality of the cell line with
continued passaging over time. Figure 49. 813 cells over-express several oncogenic proteins. General characteristics of 813 Protein was isolated from whole cell lysate of the indicated cell lines and used to western blot for the indicated proteins. cell line Murine Th1 (mTH1) and Th2 (mTH2) cells were used as normal controls. CTLL-2 and HUT78 are mouse and human T-cell lymphoma cell lines respectively. Jurkat cells are a transformed Cytogenetic analysis revealed that T-cell line and Jeko cells are a B-cell MCL cell line used as a positive control. Early and late passages of 813 cells the 813 cell line has an unstable overexpress PRMT5, cyclin D1, cyclin D3, and symmetric dimethyl-H4R3 (H4R3Me2S) relative to normal murine Th1 and Th2 cells. There is no difference in cyclin E and p65 expression genomic signature. Trisomy of in 813 cells relative to Th1 and Th2 cells. chromosome 15 was common to all of the clones examined from the early passage of the cell line and 2 of the 3 clones contained deletions in chromosome 6 (Figure 48A). The late passage of the cell line displayed even more abnormalities. Four different clones were examined from the late passage of the cell line and except one were tetra and hexaploid. A representative picture from one of the clones (Figure 48B) demonstrated two translocations (15;16) and
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Figure 50. 813 cells are sensitive to PRMT5 inhibition with CMP5.
A) 813 cells treated with CMP5 displayed a decrease in cell viability measured by annexin V and PI flow cytometry and B) a decrease in cellular proliferation as measured by MTS assay.
(4;23). Thus both passages of the cell line have an unstable genomic signature which increases in instability as the cell line is passaged.
In order to delve deeper into the characteristics of the 813 cell line the baseline expression of proteins known to be commonly overexpressed in cancer were analyzed by western blotting whole cell lysate from both early and late passages of the cell line
(Figure 49). Murine Th1 (mTh1) and Th2 (mTh2) cells were used as negative controls for the panel. The mouse T-cell lymphoma cell line, CTLL-2, and the human T-cell lymphoma cell line, Hut-78, were included as positive controls. Jeko, a MCL cell line shown previously to overexpress PRMT5, and Jurkat cells, also previously shown to express
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Figure 51. CMP5 treatment in 813 cells results in a loss of cyclin D1 and c-myc.
Late passage 813 cells were treated with CMP5 and analyzed for the indicated protein expression via western blot analysis. A) Inhibition of PRMT5 activity with CMP5 treatment resulted in a loss in cyclin D1 protein expression. B) Late passage 813 cells were treated with 25 µM CMP5 and protein was collected at the indicated timepoints. C-myc protein expression was loss within 1 hour of CMP5 treatment.
many of the proteins analyzed in the panel, were also included as controls. Both the early and late passages of 813 cells overexpress PRMT5 and one of its epigenetic marks symmetric dimethyl-H4R3, relative to normal murine Th1 and Th2 cells. The cells also overexpressed cyclin D3 relative to the normal controls. Cyclin D1 and c-myc are known to be dysregulated in various types of lymphoma and to cooperate with PRMT5 to increase tumorigenesis. Not surprisingly both passages of the 813 cell line overexpressed cyclin D1 and c-myc albeit at a lower level than the positive control, Jeko cells.
Use of a PRMT5 small molecule inhibitor in 813 cells causes changes in activation and cytokine signaling
We surmised that since the Tg813 cell line was derived from a hPRMT5 transgenic mouse it would be dependent on PRMT5 for its growth and proliferation. The small molecule inhibitor CMP5 (described in the previous chapter) was titrated in 813 cells and as expected, the inhibition of PRMT5 activity with CMP5 in 813 cells resulted in apoptosis
(Figure 50A) and a decrease in proliferation (Figure 50B). Treatment of 813 cells with
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Figure 52. PRMT5 modestly regulates T-cell activation 813 cells.
Early and late passages of 813 cells were activated with plate-bound anti-CD3 and soluble CD28 either co- current with CMP5 treatment (co-treat) or for 24 hours followed by CMP5 treatment (pre-activation). Cells were collected 24 and 48 hours after CMP5 treatment and analyzed by flow cytometry for CD25 and CD69 T- cell activation markers. A) Early and late passage cells did not express CD25 or CD69 when unactivated. B) Early passage cells activated and robustly produced CD69 as seen in the DMSO control. Co-treatment with CMP5 moderately blunted the activation response while CMP5 had little to no effect in the pre-activation group. C) The activation response in late passage cells was blunted as compared to the early passage cells. However, similar to the early passage cells CMP5 co-treatment increased the production of CD69 but displayed less of an effect in the pre-activated cells. Neither of the cell lines expressed appreciable amounts of CD25 with or without activation. Continued on next page.
148
Figure 52. Continued from previous page.
the LC50 dose of CMP5 determined from the previous results resulted in a reduction in cyclin D1 protein (Figure 51A). CMP5 treatment also resulted in the time dependent loss of c-myc protein which occurred within 2 hours after treatment in the cells (Figure 51B).
These results suggest that PRMT5 drives either the over-expression of or maintains the stability of these oncogenic proteins in this cell line and that blockade of PRMT5 can attenuate these malignant effects. Since 813 cells were determined to be a T-cell line, we wanted to determine if they could be activated in a manner similar to normal T-cells. 813 cells were activated with plate bound anti-CD3 antibody and soluble CD28. The cells were activated under two conditions: either activation with co-treatment with CMP5 or
149
Figure 53. 813 cells produce more IL-4 than IFN-γ.
Ai) Early and late 813 cells were treated with CMP5 and supernatant was collected and analyzed for IL-4 or INF-γ expression by ELISA. Aii) Early and late 813 cells were also activated with plate bound anti-CD3 and soluble CD-28 and either treated with CMP5 concurrently (co) or pre-activation for 24 hours followed by CMP5 treatment (pre). Early passages produce slightly more IL-4 and INF-γ than late passage cells Ai) IL-4 production is attenuated by PRMT5 inhibition with CMP5 treatment in both early and late passages of the 813 cell line however attenuation is more dramatic in the early passage cells. T-cell activation greatly increases IL-4 production in the early passage cells but only modestly so in the later cell passage. Aii) T-cell activation and simultaneous co-treatment of 813 cells with CMP5 attenuated IL-4 production in both early and late 813 cell passages, the effect of which is most dramatic at the 48 hour timepoint. Pre-activation for 24 hours followed by CMP5 treatment had the reverse effect in early passage and no effect in late passages of the cell line. Bi) The cells at baseline produced less IFN-γ than IL-4 and CMP5 treatment had little to no effect on IFN-γ production in either passage of the cell line. Bii) T-cell activation increased IFN-γ production in the early passage 813 cells but not the later passage. CMP5 treatment in either the co-treated or pre- activated group for the early passage cells resulted in blunted INF-γ production. 150
pre-activation for 24 hours followed by
treatment with CMP5. Cells were analyzed
by flow cytometry for the expression the
T-cell activation marker CD69 and CD25, a
component of the IL-2 receptor that
becomes expressed when T-cells are
activated. As anticipated, the expression of
the T cell activation marker CD69 was
stimulated upon activation, although the
early passage cells activated more robustly Figure 54. SCID mice engrafted with 813 cells develop an aggressive T-cell lymphoma. than the late passage cells as can be seen Representative immunohistochemistry of SCID mouse engrafted with 100,000 813 cells. The formalin-fixed by comparing the non-activated cells with paraffin-embedded spleen from the engrafted mouse was stained with hematoxylin and eosin in addition to antibodies against the B-cell marker CD20, the T-cell the DMSO control (Figure 52A, Figure markers CD3, CD4 and CD8, PRMT5, the hematopoietic progenitor cell marker CD34, the pre- 52B, and Figure 52C). Surprisingly lymphoid marker TdT, the T-cell migration and adhesion marker CD99 and the proliferation marker inhibition of the methyltransferase activity Ki-67. Engrafted SCID mice develop a rapidly proliferating T-cell lymphoma that highly expresses CD3, CD4, PRMT5, TdT and Ki67. of PRMT5 resulted in a slight increase in
CD69 expression with activation seeming to suggest that PRMT5 may aid in the regulation of cytokine production in the cell line.
Due to the high expression of surface CD4 on the 813 cells it was theorized that the cells closely resembled T helper cells. Th1 cells typically produce IFN-γ while Th2 cells typically produce IL-4 thus an ELISA assay was used to analyze the IFN-γ and IL-4 levels in early
151
B
Figure 55. 813 cells cause disease in SCID mice.
A) Survival curve of SCID mice (5 per group) engrafted with various concentrations of 813 cells. Mice were either engrafted with 1,000,000, 500,000, 250,000, or 100,000 cells via tail vein injection. All engrafted mice had succumbed to disease by 24 days. B) The immunophenotype of splenocytes isolated from SCID mice after they were engrafted with 813 cells and developed disease was analyzed by flow cytometry. The splenocytes from these mice expressed CD3, CD4 and CD5 similar to the 813 cells.
and late passage cells under various conditions. Early passages of the cells produced modestly more IL-4 than the later passages as can be seen by the bars for the DMSO control condition for both cell lines (Figure 53 A). The production of IL-4 could be attenuated by inhibition of the methyltransferase activity of PRMT5 with use of the small molecule inhibitor CMP5 (Figure 53Ai). T-cell activation was carried out either concurrent with CMP5 treatment or 24 hours prior to CMP5 treatment as described previously. T-cell activation greatly increased IL-4 production in the early passage of 813 cells but only modestly increased in late 813 as compared to the DMSO control (Figure
53Aii). Additionally, pre-activation in early 813 cells followed by CMP5 treatment seemed to increase IL-4 production. The 813 cells produced substantially less IFN-γ than
152
IL-4. T-cell activation had little effect
on IFN-γ production in late passage
cells but increased production,
though not to the same extent as IL-
4 production, in early passage cells
Figure 56. 813 cells cause disease in FVB/N mice. (Figure 53Bii). Moreover, CMP5
Survival curve of FVB/N mice (4 per group) engrafted with 813 cells. Mice were either engrafted with 1,000,000, treatment also had little to no effect 5,000,000, 10,000,000, or 20,000,000 cells via tail vein injection. All engrafted mice regardless of the amount of cells on the IFN-γ production in late engrafted had succumbed to disease by 22 days. passage cells while production was blunted in early passage cells as seen previously with IL-4 (Figure 53Bi). These experiments seem to suggest that PRMT5 helps to drive cytokine production in these cells. Since the cells produce more of the cytokine IL-4 than IFN-γ it appears that they may be Th-2 cells, however, more experiments will have to be conducted to confirm a definitive phenotype.
813 cells cause malignant tumors in mice
We also wanted to evaluate the malignant potential of the 813 cell line therefore we engrafted 813 into severe combined immunodeficiency (SCID) mice. Different concentrations of late passage 813 cells ranging from 100,000 to 1,000,000 were engrafted via tail vein injection into 7 week old SCID mice. Within 23 days all of the mice had succumbed to disease characterized by splenomegaly, hepatomegaly and nephromegaly and localized solid tumors in these organs (Figure 55A). Splenic tissue
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Figure 57. 813 cells cause lymphoma in FVB/N mice.
A) Engrafted mice presented with hepatomegaly, nephromegaly and splenomegaly relative to wild type mice of the same age. B) The average weights of the spleen, liver and kidneys of engrafted mice were significantly higher than the average weights of unengrafted control mice. Organs from four mice were used for the kidney and liver values while 8 mice were used for the spleen values. *p<0.001
from engrafted mice demonstrated a high expression of PRMT5, CD3, TdT concurrent with high Ki-67 expression and moderate CD4 expression (Figure 54). Additionally, flow cytometry analysis revealed that splenocytes isolated from these mice possessed an immunophenotype similar to the engrafted 813 cells and expressed high surface CD4 and CD43 and a moderate amount of surface CD5 and CD3 (Figure 55B). Next we wanted to examine the effects of 813 cells in a syngeneic model, they were engrafted into immune competent FVB/N mice. Various concentrations of cells from 1 to 20 million were engrafted via tail vein injection as performed previously. Similar to the results from the SCID mice, the FVB/N mice succumbed to disease within 21 days (Figure 56). Also similar to the SCID mice, FVB/N mice presented with significant splenomegaly, nephromegaly, and hepatomegaly relative to wild type unengrafted FVB/N mice (Figure
57A and Figure 57B). The mice also presented with solid tumors most often in the liver 154
Figure 58. Immunophenotype of splenocytes from control and 813 engrafted mice.
Splenocytes were isolated from wild type (wild type control) FVB/N mice and mice engrafted with the 813 cell lines and analyzed for surface markers by flow cytometry. Engrafted mice expressed higher amounts of surface B and T lymphocyte markers as examined by flow cytometry. Most notably, the expression of the T- cell markers CD3, CD4 and CD5 more than doubled from control to engrafted mice expression.
and kidneys. Immunophenotypically, splenocytes from engrafted mice expressed more surface CD4, CD43, CD5, CD3 and to a lesser extent CD19 and B220 relative to unengrafted mice (Figure 58).Protein isolated from splenocytes of the 813 engrafted mice over expressed PRMT5 and its methylation mark symmetric (2Me) H4R3 relative to the unengrafted mice (Figure 59).
Deletion of PRMT5 decrease cellular proliferation in 813 cells
In order to assess how important PRMT5 was to the malignancy of the cells we used an inducible Crispr-Cas9 gene editing system to knockout PRMT5 expression. Two separate single guide RNAs, induced to excise PRMT5 with doxycycline, were able to completely
155
Figure 59. FVB/N mice engrafted with 813 cells express oncogenic proteins.
Protein lysates from splenocytes isolated from wild type (control) and 813 engrafted FVB/N mice were analyzed by western blotting for PRMT5, c-myc and sym-H4R3(2me) expression. Control FVB/N mice express very little PRMT5 in contrast to mice engrafted with 813 cells which highly express PRMT5. In general engrafted mice over-expressed c-myc and sym-H4R3(2Me) relative to control mice. Primary tumor cells from one of the engrafted mice (#72T), 813 cells, Jurkat, the MCL cell line Jeko and the murine MCL cell line FC- mMCL were used as positive controls.
obliterate PRMT5 protein expression in 813 cells (Figure 60A). Loss of PRMT5 expression decreased the absolute number of cells as compared to control cells (Figure 60B), indicating that PRMT5 is important to the cellular proliferation of the 813 cell line. These experiments highlight the highly malignant nature of the 813 cell line in mice, and present an opportunity to use this cell line as an in vivo model to study T-cell lymphoma as well as the role of PRMT5 in this disease.
Discussion
PRMT5 is an important oncogene that has been linked to a number of malignancies such as various subtypes of NHL, including mantle cell lymphoma which was presented in chapter 2. Since most cases of NHL occur in B-cells, and PRMT5 has been studied the most extensively in the context of B-cell lymphoma we created a transgenic mouse that utilized the Eµ enhancer/promoter to drive hPRMT5 overexpression in the B-cells of 156
Figure 60. Inducible deletion of PRMT5 in 813 cells results in decreased proliferation.
Tg813 cells were transduced with a lentivirus containing Cas9 to create cells that constitutively express Cas9. Cells were then transduced with a lentivirus with blasticidin-resistant, GFP-tagged single guide RNA targeted to PRMT5. A) Treatment with doxycycline induced the knockout of PRMT5 and protein expression was loss within 24hours. B) Loss of PRMT5 protein expression correlated with decreased cellular proliferation.
FVB/N mice. We hypothesized that PRMT5 was a sole driver of B-cell lymphomagenesis and that over-expression of hPRMT5 in the B-lymphocytes of a mouse would increase incidence of lymphoma in these mice. Although hPRMT5 mice developed lymphoma at a higher rate than non-transgenic mice the model still had a lower than expected penetrance. One explanation for this low penetrance could be the existence of mosaicism in some of the founders. Indeed up to 30% of transgenic mice can display
157 some form of mosaicism [238]. The best example of mosaicism in our model would be the line of founder 15 in which less than 10% of the progeny expressed the transgene.
The low penetrance of the model may also in part be due to the cooperative nature of
PRMT5. Indeed previous studies have demonstrated that PRMT5 cooperates with a number of other chromatin remodelers and oncogenes in order to promote transformation [175,176,237]. Thus in order to improve the penetrance of the model it may be necessary to cross the hPRMT5 transgenic mouse with other oncogenic strains to improve the penetrance and phenotype of the model. Some potential strains include the
Eµ-myc mouse, which overexpresses myc or the Eμ-D1T286A transgenic mouse, which constitutively expresses nuclear cyclin D1, both of which are targeted to the B-cell lymphoid compartment by the immunoglobulin enhancer/promoter [239,240]. Overall the hPRMT5 transgenic mouse highlights the role of PRMT5 as a driver of transformation and lymphomagenesis but also points to the necessity of the dysregulation of other oncogenes in order to fully potentiate a malignant phenotype.
In hPRMT5 transgenic mice, PRMT5 mRNA was largely over-expressed in the B-cell lymphocyte compartment and to a lesser extent in the NK- and T-cell compartments.
Thus hPRMT5 transgenic mice developed mostly B-cell lymphomas characterized by organomegaly, concurrent with lymphocyte infiltrate in these and other organs.
Interestingly, a number of the lymphomas generated from the hPRMT5 were determined to be T-cell lymphomas. In fact we were also able to isolate tumor cells and generate a T- cell lymphoma cell line (Tg813) from a tumor of a transgenic mouse. This is not
158 completely surprising since Eµ has been shown to drive gene expression in both B- and
T-cells [241]. Additionally, injected 5-fluorouracil (5-FU)–treated bone marrow hematopoietic stem/progenitor cells (HSPC) transduced with retroviral supernatants encoding PRMT5 and cyclin D1T286A caused mature T-cell lymphomas in lethally irradiated, syngeneic C57BL/6 mice [175]. It is unclear why the presence of PRMT5 in the lymphocyte compartment might preferentially select for T-cell malignancies, but previously published studies may offer insight into the role of PRMT5 in normal T-cell biology and explain why PRMT5 dysregulation results in these malignancies. Cytokine gene transcription in T helper (Th) cells is regulated by arginine methylation of the NFAT cofactor NIP45 [242]. Additionally, another study identified PRMT5 as an important factor that promoted Nuclear Factor of Activated T-cells (NFAT)-driven promoter activity and
IL-2 secretion in T-cells [147]. Furthermore loss of PRMT5 in hematopoietic stem cells results in a significant decrease in thymus size and T-cell cellularity without an apparent change in spleen size or cellularity or the number of B-cells in this compartment [146].
Thus potentially PRMT5 exerts more control over T-cell biology and PRMT5 dysregulation alone in the T-cell compartment is able to promote malignancy, whereas
B-cell dysregulation, due to unknown reasons, requires other oncogenic factors besides
PRMT5.
The 813 cell line is a mature T-cell line that grows without the need for external cytokines and expresses surface CD43, CD45, CD24, CD4 CD5 and CD3 in early passages. Later passages lose CD3 surface expression but continue to express intracellular CD3. Earlier
159 passages are also polyclonal for the TCR Vβ receptor while later passages of the cell line are monoclonal for the Vβ-17 subunit. Although we do not have a definite explanation for this phenomenon, loss of surface CD3 expression may be tied to the instability of the cell line. Indeed cytogenetic analysis revealed a number of insertions, deletions, incidences of trisomy and chromosomal profile that evolves from diploid/triploid to tetraploid/hexaploid with serial passaging. Alternatively, it could be the result of a single aggressive clone that expanded and took over the cell culture with subsequent passages.
813 cells over-expressed PRMT5 and its epigenetic mark symmetric dimethyl-H4R3 as well as classical oncogenic proteins including cyclin D1, D3 and c-myc. Furthermore we were able to show that inhibition of PRMT5 with the small molecule inhibitor CMP5 attenuated cyclin D1 and c-myc expression in 813 cells.
At this time it is unclear exactly what type of T-cell lymphoma the 813 cell line represents however cursory ELISA analysis determined that the cells produced IL-4 and little to no
IFN-γ. Additionally, the cells could be activated with methods similar to normal T-cells.
We also observed modulation in both cytokine production and activation with PRMT5 inhibition indicating that PRMT5 plays a major role in these functions in this cell line.
Ultimately more in-depth studies such as RNA sequencing and/or comprehensive cytokine array analysis may shed more light on the phenotype of the cell line and allow for a definitive characterization. Despite not having a definitive phenotype for the cells, we engrafted the cell line into two different strains of mice to test for its ability to cause disease. SCID mice engrafted by tail vein injection with 813 cells developed hepatic, renal
160 and/or splenic tumors with as few as 100,000 cells and succumbed to disease within 20 days. More impressively, fully immune competent FVB/N mice engrafted with 813 cells also developed hepatic, renal and/or splenic tumors with a few as 1,000,000 cells. Based on this data we are confident that we have developed a murine syngeneic model of lymphoma that potentially can be used to study the role of PRMT5 in a lymphoma microenvironment and in the context of a normal immune system. This is important because although T-cell lymphomas represent only 15% of NHLs the overall mortality rate of T-cell NHLs is higher than that of B-cell NHLs as a whole due largely in part to a lack of knowledge about the mechanism of disease because of its rarity [243]. Thus perhaps our model can shed light on mechanisms of T-cell lymphomagenesis regulated by PRMT5 as well as allow researchers to utilize a mouse model to test potential therapeutic agents.
In conclusion, this chapter describes the successful creation of the first known PRMT5 overexpression transgenic mouse model. Despite the low penetrance of the model it can be used to evaluate the role of PRMT5 in lymphomagenesis in the context of other oncogenic drivers. This model also yielded the T-cell lymphoma Tg813 cell line which causes a highly malignant lymphoma when engrafted into mice. Both of these models prove that PRMT5 is an important player in lymphomagenesis and both can used to study more in depth the role of PRMT5 in both B-and T-cell lymphoma.
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Chapter 4: Concluding Discussion
Summary
Epigenetics, or the heritable, post-transcriptional changes in gene expression, is responsible for a number of normal physiological processes as well as the development of cancer. One such epigenetic modifier, protein arginine methyltransferase 5 (PRMT5), is important in embryogenesis, hematopoiesis, adipogenesis, and keratinocyte homeostasis
[133,146,149,151]. However, despite regulating a number of normal physiological processes dysregulation of PRMT5 has also been implicated in many malignancies including glioblastoma multiforme, breast cancer, hepatocellular carcinoma, testicular cancer, and leukemias and lymphomas [26,155,157,158,164,230]. The discovery of the relevance of PRMT5 in lymphoma, and non-Hodgkin’s lymphoma in particular, offered an opportunity to pursue a potential therapeutic target in a disease cohort in need of new curative remedies. Specifically in lymphomas, PRMT5 appears to be a master regulator that cooperates with other commonly dysregulated oncogenes such as cyclin
D1, c-myc and PRC2 members promoting cell cycle abnormalities and unmediated proliferation [153,155,175,237]. Additionally, PRMT5 becomes over-expressed in B-cells undergoing Epstein-Barr virus-mediated transformation and B-cell transformation is prevented with PRMT5 inhibition [199]. Based on these data we hypothesized that
162
PRMT5 is a major driver of B-cell transformation in lymphomagenesis. To test our hypothesis we first aimed to characterize a first in its class PRMT5 small molecule inhibitor in a B-cell lymphoma malignancy and second we aimed to characterize a first described PRMT5 transgenic mouse. The overall goal of this project was to not only establish PRMT5 as a major driver of lymphoma but to glean more in-depth information about the transformative effects of PRMT5 in the context of the disease that can be used to ultimately create more potent therapeutics for the clinic.
In order to test our hypothesis in our first aim we chose the B-cell NHL subset MCL as a disease model. Previously published data indicated that PRMT5 was an important component of the malignant nature of MCL by demonstrating that PRMT5 is overexpressed in MCL relative to normal, resting B-cells and that re-expression of repressed miRs attenuated PRMT5 expression and de-repression of tumor suppressor miRs concurrent with cell death [127]. Building upon the previous study we were able to confirm that PRMT5 was highly over-expressed in primary MCL patient samples as well as in MCL patient-derived cell lines as compared to resting normal B-cells. Furthermore we were able to show that increased nuclear expression of PRMT5 directly correlated with more aggressive variants of the disease. It is worth noting that in some malignancies nuclear PRMT5 is associated with more malignant and aggressive disease while in others cytoplasmic PRMT5 is associated with more malignant and aggressive disease. There is no clear explanation for this phenomenon, however, oncogenic factors in some diseases may require the methyltransferase activity of PRMT5 in the nucleus to
163 turn off important tumor suppressor genes, while other diseases may need PRMT5 in the cytoplasm to silence cell cycle regulatory proteins or cooperate with other oncogenic proteins. Notably PRMT5 can interact with the SWI/SNF chromatin remodeling complex to silence genes [125]and can cooperate with cyclin D1 to silence p53 [244]. Due to the large variety of oncogenic complexes and proteins that interact with PRMT5, it is difficult to surmise which set of complexes may be in play in MCL.
Additionally, this body of work is the first to describe the action of PRMT5 small molecular inhibitors, first described in our lab [199], in the context of MCL. As mentioned previously, the majority of MCL clinical cases overexpress cyclin D1 due to the t(11;14)(q13;q32) translocation thus dysregulation of the cell cycle is a hallmark in this disease. Use of a PRMT5 inhibitor in MCL restored the expression of cell cycle regulatory proteins and genes and decreased cellular proliferation and viability demonstrating that
PRMT5 is intricately tied to abnormal cell cycle regulation in MCL. Notably, PRMT5 was found to directly interact with cyclin D1 and control both cyclin D1 and c-myc protein. At this time it is unclear what the relationship is between cyclin D1 and PRMT5 and how or why they form a complex with one another in MCL, nevertheless, use of PRMT5 inhibitors potentially can be used to help describe this relationship in depth. Recently published work indicated that mutated cyclin D1, such as that which occurs in MCL, can promote the interaction between PRMT5 and MEP50 enhancing their methyltransferase activity and promoting malignancy [175]. Based on these data a logical relationship between cyclin D1 and PRMT5 in MCL emerges whereby the cyclin D1 translocation occurs,
164 producing a more stable cyclin D1 that is refractory to nuclear exportation and subsequent degradation. Cyclin D1 in turn phosphorylates MEP50 to encourage the binding with PRMT5 and increase their methyltransferase activity which drives lymphomagenesis by disabling key tumor suppressors such as ST7 and p53. Potential future experiments include exploring the localization and the methylation status of cyclin
D1 with and without inhibitor to determine if inhibition of the methyltransferase activity of PRMT5 affects the localization and/or stability of cyclin D1 in MCL. Additionally, site- directed mutagenesis could be used to obtain a clearer picture of how and where cyclin
D1 and PRMT5 interact with each other. The results of these studies could provide valuable insight into the pathogenesis of PRMT5, particularly in MCL, and into how
PRMT5 exploits other oncogenic proteins to drive lymphomagenesis.
This work also sheds light on PRMT5-mediated repression of tumor suppressors and cell cycle regulators. Treatment with CMP5 in MCL cell lines resulted in the de-repression of
ST7, confirming a relationship between the two proteins previously published [130,245].
We also discovered that inhibition of the methyltransferase activity of PRMT5 increased the expression of the transcription factor C/EBPβ which has also been implicated in the regulation of the cell cycle, as well as the de-repression of the transcription factor PDCD4 which controls myc [232]. This reveals a picture of PRMT5 as a master regulator that not only drives the expression of oncogenes and oncogenic proteins but represses cell cycle and survival regulators effectively solely selecting for a malignant phenotype.
165
Currently, we have begun preliminary work with 4th generation PRMT5 inhibitors that have activity in the low micromolar range and thus are more potent than CMP5. These inhibitors were created by using modeling software to add different functional groups onto the CMP5 backbone and exploring their effects on docking the molecule into the
SAM binding site in the published structure of hPRMT5. Additionally, as mentioned previously, the end goal of this project is to develop inhibitors that can be used in the clinic however a huge obstacle to achieving this goal has been a lack of in vivo data due to severe toxicity as our current inhibitors and delivery solvent system is incompatible with viability in murine models of lymphoma. CMP5 is only soluble in DMSO and intraperitoneal injection of the drug causes major gastrointestinal toxicity. Thus in addition to creating more potent inhibitors we must develop a delivery system that allows our drug to reach therapeutic levels in the blood without causing organ toxicity.
Potential drug delivery alternatives include nanoparticles or transdermal patches
[246,247].
Despite the current in vivo hurdles, in vitro data with the PRMT5 inhibitor in conjunction with other chemotherapeutic epigenetic inhibitors was very promising. Strikingly, sublethal doses of CMP5 combined with the HDAC inhibitor TSA and the DNMT inhibitor
5-aza, was as toxic or more toxic to the cells compared to single agent CMP5 treatment.
Since PRMT5 is responsible for many physiological processes including hematopoiesis, muscle stem cell renewal, lipogenesis, sodium channel generation in the heart and keratinocyte proliferation [144,146,151,170,248] we can expect that inhibition of PRMT5
166 may result in side effects such as loss of lymphoid cellularity, muscle weakness, cardiac muscle toxicity and skin disorders. Therefore, using lower doses of CMP5 with other therapeutic agents and in particular epigenetic modifiers, may alleviate the risk of side effects while still maintaining therapeutic efficacy.
Our second aim was to create a PRMT5 transgenic mouse and in the third chapter of this dissertation we described the first PRMT5 transgenic mouse. In an effort to recapitulate
PRMT5-driven B-cell NHL we used the immunoglobulin heavy chain enhancer/promoter,
Eµ, in combination with a vector containing hPRMT5 to drive the over-expression of hPRMT5 in murine B-lymphocytes. From this model we obtained 5 hPRMT5 founder mice that contained a high copy number of PRMT5 as detected by PCR, and all except founder 15 were able to pass the over-expression of PRMT5 on to progeny. Although we anticipated that hPRMT5 transgenic mice would develop B-cell lymphoma at a high rate, the model had a lower than expected incidence rate of lymphoma. As discussed previously, we believe the penetrance of the model was lower than normal due to mosaicism or the lack of complete integration of the transgene into the murine genome, preventing the transmission of the gene to germ cells and subsequently to progeny.
Additionally, the penetrance of the model could potentially be increased by crossing hPRMT5 transgenic mice with other oncogenic transgenic mice. Despite limitations in the model we were able to obtain several B-cell lymphomas which were characterized by splenomegaly, hepatomegaly, nephromegaly and increased expression of surface IgM,
CD19 and B220 in unpurified splenocytes.
167
Moreover, we discovered that PRMT5 was not only over-expressed in B-cells but in NK- and T-cells as well. Indeed, we also obtained a small number of T-cell lymphomas from this model as well including the mouse that led to the creation of the 813 cell line. While unexpected, it is not uncommon for the Eµ enhancer/promoter to be active in other lymphocyte compartments, particularly T-cells. As such the Eµ-cyclin D1 mouse also generates T-cell lymphomas [249]. T-cell lymphoma is exceedingly rare in Western countries representing 5 – 10% of all NHLs in contrast to Asian countries where it represents 15 – 20% [250]. As compared to B-cell NHLs, T-cell NHL is even more heterogeneous making it an exceedingly difficult disease to treat. The scarcity of models coupled with the rarity of the disease also makes T-cell NHL a difficult malignancy to study for the development of therapeutics. Thus the Tg813 cell line developed from our transgenic mouse could potentially be used as a tool to model T-cell lymphoma in an effort to learn more about the disease and potential therapeutic pathways.
Furthermore, the hPRMT5 gene was randomly inserted into the murine genome thus we cannot rule out the role of hPRMT5 in the development of other cancer types in this model. This is evidenced by a small cohort of adenocarcinomas, mostly of lung origin, that were also identified in the hPRMT5 transgenic mouse. However, the FVB/N strain is particularly susceptible to developing solid tumors such as hepatic and lung carcinomas
[251] thus it is plausible that these tumors were unrelated to the presence of hPRMT5.
168
Limitations of the study
Despite the promising results outlined thus far this body of work includes a few limitations. One major limitation is the lack of confirmatory experiments with other methods of PRMT5 knockdown to compare to our small molecule inhibitor. As presented in chapter 2 si/shRNA is an inefficient method of knockdown in our suspension cells, however, during the preparation of this dissertation another inhibitor was published that could be used to compare our results. Additionally, other methods of gene knockdown including the Crispr-Cas9 system could be more effective and should be explored.
Another limitation to our work is the high LC50 for our lead compound, CMP5. Currently the LC50 value, upwards of 50 µM in MCL cell lines, is prohibitive for the clinical development of the drug. Thus there is a need for us to work closely with the drug development team to design and create more potent inhibitors that are just as effective as CMP5 but with lower LC50s. Additionally, prior to clinical development we need to be able to demonstrate efficacy of the drug in an in vivo system which we currently have been unable to do. As mentioned previously, injection of the drug into mice causes severe gastrointestinal toxicity. This potentially is caused by the drug itself or the solvent and delivery system of the drug. Therefore future drug design will need to take into account alternative drug formulations as well as routes of administration to optimize delivery while minimizing toxicity.
In the third chapter of this dissertation we described the first of its kind PRMT5 transgenic mouse. Despite careful consideration to design of the model we did not
169 obtain a high incidence of disease in this model potentially due to reasons previously mentioned. Unfortunately the colony was lost therefore we will be unable to explore methods to increase the penetrance of the model. Aside from the hPRMT5 transgenic mouse, we also successfully developed a T-cell lymphoma cell line from one of the mouse tumors. Although we have made significant strides in describing the cell line, many gaps still currently exist in our characterization, thus we do not have a definitive lymphoma type for the Tg813 cell line.
Future directions
This dissertation presents many opportunities for continued exploration of the role of
PRMT5 in lymphoma. We have recently obtained 4th generation PRMT5 inhibitors from our drug design collaborators that we hope to test in MCL cell lines. Preliminary analysis has revealed that these inhibitors actively block PRMT5 methyltransferase activity and promote oncogenic cell death in the low micromolar range. Experiments such as those laid out in chapter 2 will need to be conducted on this new batch of inhibitors to determine their feasibility for in vivo studies.
We obtained promising results from the hPRMT5 transgenic mouse however, we lost the original colony. We are currently in the process of re-establishing the colony using the
C57bl/6 strain which is a more commonly used strain and will give us more options for crossing with other transgenic mouse models to increase penetrance. In addition to re- creating and characterizing the hPRMT5 transgenic mouse, further studies need to be carried out on the Tg813 cell line. Ideally, we would like to conclusively determine the
170 type of lymphoma this cell line represents. Potential experiments to be completed include cytokine array and RNA and/or DNA sequencing analysis in addition to more immunophenotyping. Overall we are confident that we have established a role for
PRMT5 in lymphomagenesis and developed tools and models that will allow for many avenues of future exploration.
171
References
1. Hamm CA, Costa FF. Epigenomes as Therapeutic Targets. Pharmacol. Ther. (2015).
2. Kohda T. Effects of embryonic manipulation and epigenetics. J. Hum. Genet. 58(7), 416–20 (2013).
3. Schäfer V, Ernst J, Rinke J, et al. EZH2 mutations and promoter hypermethylation in childhood acute lymphoblastic leukemia. J. Cancer Res. Clin. Oncol. 142(7), 1641–1650 (2016).
4. Robaina MC, Mazzoccoli L, Arruda VO, et al. Deregulation of DNMT1, DNMT3B and miR-29s in Burkitt lymphoma suggests novel contribution for disease pathogenesis. Exp. Mol. Pathol. 98(2), 200–207 (2015).
5. KIT S. Deoxyribonucleic Acids. Annu. Rev. Biochem. 32, 43–82 (1963).
6. Jones PA. The Role of DNA Methylation in Mammalian Epigenetics. Science (80-. ). 293(5532), 1068–1070 (2001).
7. Jaenisch R, Bird A. Epigenetic regulation of gene expression: how the genome integrates intrinsic and environmental signals. .
8. Du J, Johnson LM, Jacobsen SE, Patel DJ. DNA methylation pathways and their crosstalk with histone methylation. Nat. Rev. Mol. Cell Biol. 16(9), 519–532 (2015).
9. Jair K-W, Bachman KE, Suzuki H, et al. De novo CpG island methylation in human cancer cells. Cancer Res. 66(2), 682–92 (2006).
10. Okano M, Bell DW, Haber DA, Li E. DNA Methyltransferases Dnmt3a and Dnmt3b 172
Are Essential for De Novo Methylation and Mammalian Development. Cell. 99(3), 247–257 (1999).
11. Holz-Schietinger C, Matje DM, Harrison MF, Reich NO. Oligomerization of DNMT3A controls the mechanism of de novo DNA methylation. J. Biol. Chem. 286(48), 41479–88 (2011).
12. Bogdanović O, Veenstra GJC. DNA methylation and methyl-CpG binding proteins: developmental requirements and function. Chromosoma. 118(5), 549–65 (2009).
13. Hamidi T, Singh AK, Chen T. Genetic alterations of DNA methylation machinery in human diseases. Epigenomics. 7(2), 247–65 (2015).
14. Brocato J, Costa M. Basic mechanics of DNA methylation and the unique landscape of the DNA methylome in metal-induced carcinogenesis. Crit. Rev. Toxicol. 43(6), 493–514 (2013).
15. Wu H, Zhang Y. Reversing DNA methylation: mechanisms, genomics, and biological functions. Cell. 156(1-2), 45–68 (2014).
16. Choy M-K, Movassagh M, Goh H-G, Bennett MR, Down TA, Foo RSY. Genome- wide conserved consensus transcription factor binding motifs are hyper- methylated. BMC Genomics. 11(1), 519 (2010).
17. Esteller M. Epigenetics in Cancer [Internet]. N. Engl. J. Med. , 1148–1159 (2008). Available from: papers2://publication/uuid/CD7DA39A-C84C-45A9-BE69- ACA42CC6625E.
18. Deaton AM, Bird A. CpG islands and the regulation of transcription. Genes Dev. 25(10), 1010–22 (2011).
19. Hodges E, Smith AD, Kendall J, et al. High definition profiling of mammalian DNA methylation by array capture and single molecule bisulfite sequencing. Genome Res. 19(9), 1593–605 (2009).
173
20. Meissner A, Mikkelsen TS, Gu H, et al. Genome-scale DNA methylation maps of pluripotent and differentiated cells. Nature. 454(7205), 766–70 (2008).
21. Jin B, Li Y, Robertson KD. DNA methylation: superior or subordinate in the epigenetic hierarchy? Genes Cancer. 2(6), 607–17 (2011).
22. Phillips T. The Role of Methylation in Gene Expression. Nat. Educ. 1(1), 116 (2008).
23. Fabbri M, Garzon R, Cimmino A, et al. MicroRNA-29 family reverts aberrant methylation in lung cancer by targeting DNA methyltransferases 3A and 3B. Proc. Natl. Acad. Sci. U. S. A. 104(40), 15805–10 (2007).
24. Garzon R, Liu S, Fabbri M, et al. MicroRNA-29b induces global DNA hypomethylation and tumor suppressor gene reexpression in acute myeloid leukemia by targeting directly DNMT3A and 3B and indirectly DNMT1. Blood. 113(25), 6411–8 (2009).
25. Braconi C, Huang N, Patel T. MicroRNA-dependent regulation of DNA methyltransferase-1 and tumor suppressor gene expression by interleukin-6 in human malignant cholangiocytes. Hepatology. 51(3), 881–90 (2010).
26. Chen B-F, Gu S, Suen Y-K, Li L, Chan W-Y. microRNA-199a-3p, DNMT3A, and aberrant DNA methylation in testicular cancer. Epigenetics. 9(1), 119–28 (2014).
27. Duursma AM, Kedde M, Schrier M, le Sage C, Agami R. miR-148 targets human DNMT3b protein coding region. RNA. 14(5), 872–877 (2008).
28. Di Ruscio A, Ebralidze AK, Benoukraf T, et al. DNMT1-interacting RNAs block gene-specific DNA methylation. Nature. 503(7476), 371–6 (2013).
29. Esteller M. Non-coding RNAs in human disease. Nat. Rev. Genet. 12(12), 861–74 (2011).
30. Wang J, Chen J, Sen S. MicroRNA as Biomarkers and Diagnostics. J. Cell. Physiol. 231(1), 25–30 (2016). 174
31. Tuna M, Machado AS, Calin GA. Genetic and epigenetic alterations of microRNAs and implications for human cancers and other diseases. Genes, Chromosom. Cancer. 55(3), 193 – 214 (2016).
32. Hill DA. Influence of linker histone H1 on chromatin remodeling. Biochem. Cell Biol. 79(3), 317–24 (2001).
33. Ellis L, Atadja PW, Johnstone RW. Epigenetics in cancer: targeting chromatin modifications. Mol Cancer Ther. 8(6), 1409–1420 (2009).
34. Lee J-SS, Smith E, Shilatifard A. The language of histone crosstalk. Cell. 142(5), 682–685 (2010).
35. Dominski Z, Marzluff WF. Formation of the 3′ end of histone mRNA. Gene. 239(1), 1–14 (1999).
36. Albig W, Doenecke D. The human histone gene cluster at the D6S105 locus. Hum. Genet. 101(3), 284–94 (1997).
37. Marzluff WF, Gongidi P, Woods KR, Jin J, Maltais LJ. The Human and Mouse Replication-Dependent Histone Genes. Genomics. 80(5), 487–498 (2002).
38. Suto RK, Clarkson MJ, Tremethick DJ, Luger K. Crystal structure of a nucleosome core particle containing the variant histone H2A.Z. Nat. Struct. Biol. 7(12), 1121–4 (2000).
39. Biterge B, Schneider R. Histone variants: key players of chromatin. Cell Tissue Res. 356(3), 457–66 (2014).
40. Luk E, Ranjan A, Fitzgerald PC, et al. Stepwise histone replacement by SWR1 requires dual activation with histone H2A.Z and canonical nucleosome. Cell. 143(5), 725–36 (2010).
41. Papamichos-Chronakis M, Watanabe S, Rando OJ, Peterson CL. Global regulation of H2A.Z localization by the INO80 chromatin-remodeling enzyme is essential for 175
genome integrity. Cell. 144(2), 200–13 (2011).
42. Binda O, Sevilla A, LeRoy G, Lemischka IR, Garcia BA, Richard S. SETD6 monomethylates H2AZ on lysine 7 and is required for the maintenance of embryonic stem cell self-renewal. Epigenetics. (2013).
43. Li A, Yu Y, Lee S-C, Ishibashi T, Lees-Miller SP, Ausió J. Phosphorylation of histone H2A.X by DNA-dependent protein kinase is not affected by core histone acetylation, but it alters nucleosome stability and histone H1 binding. J. Biol. Chem. 285(23), 17778–88 (2010).
44. Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM. DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139. J. Biol. Chem. 273(10), 5858–5868 (1998).
45. Rasmussen TP, Huang T, Mastrangelo M-A, Loring J, Panning B, Jaenisch R. Messenger RNAs encoding mouse histone macroH2A1 isoforms are expressed at similar levels in male and female cells and result from alternative splicing. Nucleic Acids Res. 27(18), 3685–3689 (1999).
46. Chadwick BP. Histone H2A variants and the inactive X chromosome: identification of a second macroH2A variant. Hum. Mol. Genet. 10(10), 1101–1113 (2001).
47. Chakravarthy S, Luger K. The histone variant macro-H2A preferentially forms “hybrid nucleosomes”. J. Biol. Chem. 281(35), 25522–31 (2006).
48. Gaspar-Maia A, Qadeer ZA, Hasson D, et al. MacroH2A histone variants act as a barrier upon reprogramming towards pluripotency. Nat. Commun. 4, 1565 (2013).
49. Shukla MS, Syed SH, Goutte-Gattat D, et al. The docking domain of histone H2A is required for H1 binding and RSC-mediated nucleosome remodeling. Nucleic Acids Res. 39(7), 2559–70 (2011).
50. Ishibashi T, Li A, Eirín-López JM, et al. H2A.Bbd: an X-chromosome-encoded histone involved in mammalian spermiogenesis. Nucleic Acids Res. 38(6), 1780–9 (2010). 176
51. Churikov D, Siino J, Svetlova M, et al. Novel human testis-specific histone H2B encoded by the interrupted gene on the X chromosome. Genomics. 84(4), 745–56 (2004).
52. Li A, Maffey AH, Abbott WD, et al. Characterization of nucleosomes consisting of the human testis/sperm-specific histone H2B variant (hTSH2B). Biochemistry. 44(7), 2529–35 (2005).
53. Boulard M, Gautier T, Mbele GO, et al. The NH2 tail of the novel histone variant H2BFWT exhibits properties distinct from conventional H2B with respect to the assembly of mitotic chromosomes. Mol. Cell. Biol. 26(4), 1518–26 (2006).
54. Montellier E, Boussouar F, Rousseaux S, et al. Chromatin-to-nucleoprotamine transition is controlled by the histone H2B variant TH2B. Genes Dev. 27(15), 1680– 92 (2013).
55. Li M, Fang Y. Histone variants: the artists of eukaryotic chromatin. Sci. China. Life Sci. 58(3), 232–9 (2015).
56. Elsaesser SJ, Allis CD. HIRA and Daxx constitute two independent histone H3.3- containing predeposition complexes. Cold Spring Harb. Symp. Quant. Biol. 75, 27– 34 (2010).
57. Hake SB, Garcia BA, Duncan EM, et al. Expression patterns and post-translational modifications associated with mammalian histone H3 variants. J. Biol. Chem. 281(1), 559–68 (2006).
58. Chen P, Zhao J, Wang Y, et al. H3.3 actively marks enhancers and primes gene transcription via opening higher-ordered chromatin. Genes Dev. 27(19), 2109–24 (2013).
59. Jin C, Zang C, Wei G, et al. H3.3/H2A.Z double variant-containing nucleosomes mark “nucleosome-free regions” of active promoters and other regulatory regions. Nat. Genet. 41(8), 941–5 (2009).
60. Sekulic N, Bassett EA, Rogers DJ, Black BE. The structure of (CENP-A-H4)(2) reveals 177
physical features that mark centromeres. Nature. 467(7313), 347–51 (2010).
61. Foltz DR, Jansen LET, Bailey AO, et al. Centromere-specific assembly of CENP-a nucleosomes is mediated by HJURP. Cell. 137(3), 472–84 (2009).
62. Henikoff S, Furuyama T. The unconventional structure of centromeric nucleosomes. Chromosoma. 121(4), 341–52 (2012).
63. Goutte-Gattat D, Shuaib M, Ouararhni K, et al. Phosphorylation of the CENP-A amino-terminus in mitotic centromeric chromatin is required for kinetochore function. Proc. Natl. Acad. Sci. U. S. A. 110(21), 8579–84 (2013).
64. Bailey AO, Panchenko T, Sathyan KM, et al. Posttranslational modification of CENP-A influences the conformation of centromeric chromatin. Proc. Natl. Acad. Sci. U. S. A. 110(29), 11827–32 (2013).
65. Niikura Y, Kitagawa R, Ogi H, Abdulle R, Pagala V, Kitagawa K. CENP-A K124 Ubiquitylation Is Required for CENP-A Deposition at the Centromere. Dev. Cell. 32(5), 589–603 (2015).
66. Izzo A, Kamieniarz K, Schneider R. The histone H1 family: specific members, specific functions? Biol. Chem. 389(4), 333–43 (2008).
67. Cheema MS, Ausió J. The Structural Determinants behind the Epigenetic Role of Histone Variants. Genes (Basel). 6(3), 685–713 (2015).
68. Terme J-M, Sesé B, Millán-Ariño L, et al. Histone H1 variants are differentially expressed and incorporated into chromatin during differentiation and reprogramming to pluripotency. J. Biol. Chem. 286(41), 35347–57 (2011).
69. Kalashnikova AA, Winkler DD, McBryant SJ, et al. Linker histone H1.0 interacts with an extensive network of proteins found in the nucleolus. Nucleic Acids Res. 41(7), 4026–35 (2013).
70. Stoldt S, Wenzel D, Schulze E, Doenecke D, Happel N. G1 phase-dependent 178
nucleolar accumulation of human histone H1x. Biol. Cell. 99(10), 541–52 (2007).
71. Martianov I, Brancorsini S, Catena R, et al. Polar nuclear localization of H1T2, a histone H1 variant, required for spermatid elongation and DNA condensation during spermiogenesis. Proc. Natl. Acad. Sci. U. S. A. 102(8), 2808–13 (2005).
72. Yan W, Ma L, Burns KH, Matzuk MM. HILS1 is a spermatid-specific linker histone H1-like protein implicated in chromatin remodeling during mammalian spermiogenesis. Proc. Natl. Acad. Sci. U. S. A. 100(18), 10546–51 (2003).
73. Mizusawa Y, Kuji N, Tanaka Y, et al. Expression of human oocyte-specific linker histone protein and its incorporation into sperm chromatin during fertilization. Fertil. Steril. 93(4), 1134–41 (2010).
74. Jagtap P, Szabó C. Poly(ADP-ribose) polymerase and the therapeutic effects of its inhibitors. Nat. Rev. Drug Discov. 4(5), 421–40 (2005).
75. Xu Y-M, Du J-Y, Lau ATY. Posttranslational modifications of human histone H3: an update. Proteomics. 14(17-18), 2047–60 (2014).
76. Hottiger MO. Nuclear ADP-Ribosylation and Its Role in Chromatin Plasticity, Cell Differentiation, and Epigenetics. Annu. Rev. Biochem. 84, 227–63 (2015).
77. Karlberg T, Langelier M-F, Pascal JM, Schüler H. Structural biology of the writers, readers, and erasers in mono- and poly(ADP-ribose) mediated signaling. Mol. Aspects Med. 34(6), 1088–108 (2013).
78. Li N, Chen J. ADP-ribosylation: activation, recognition, and removal. Mol. Cells. 37(1), 9–16 (2014).
79. Messner S, Hottiger MO. Histone ADP-ribosylation in DNA repair, replication and transcription. Trends Cell Biol. 21(9), 534–42 (2011).
80. Sawicka A, Seiser C. Sensing core histone phosphorylation - a matter of perfect timing. Biochim. Biophys. Acta. 1839(8), 711–8 (2014). 179
81. Xiao A, Li H, Shechter D, et al. WSTF regulates the H2A.X DNA damage response via a novel tyrosine kinase activity. Nature. 457(7225), 57–62 (2009).
82. Rossetto D, Avvakumov N, Côté J. Histone phosphorylation: a chromatin modification involved in diverse nuclear events. Epigenetics. 7(10), 1098–108 (2012).
83. Sarkar P, Reichman C, Saleh T, Birge RB, Kalodimos CG. Proline cis-trans Isomerization Controls Autoinhibition of a Signaling Protein. Mol. Cell. 25(3), 413– 426 (2007).
84. Schmidpeter PAM, Jahreis G, Geitner A-J, Schmid FX. Prolyl isomerases show low sequence specificity toward the residue following the proline. Biochemistry. 50(21), 4796–803 (2011).
85. Howe FS, Mellor J. Proline cis-trans isomerization is influenced by local lysine acetylation-deacetylation. Microb. Cell. 1(11), 390–392 (2014).
86. Nelson CJ, Santos-Rosa H, Kouzarides T. Proline isomerization of histone H3 regulates lysine methylation and gene expression. Cell. 126(5), 905–16 (2006).
87. Cao J, Yan Q. Histone ubiquitination and deubiquitination in transcription, DNA damage response, and cancer. Front. Oncol. 2, 26 (2012).
88. Hochstrasser M. Ubiquitin-dependent protein degradation. Annu. Rev. Genet. 30, 405–39 (1996).
89. Weake VM, Workman JL. Histone ubiquitination: triggering gene activity. Mol. Cell. 29(6), 653–63 (2008).
90. Pinder JB, Attwood KM, Dellaire G. Reading, writing, and repair: the role of ubiquitin and the ubiquitin-like proteins in DNA damage signaling and repair. Front. Genet. 4, 45 (2013).
91. Nijman SMB, Luna-Vargas MPA, Velds A, et al. A genomic and functional inventory 180
of deubiquitinating enzymes. Cell. 123(5), 773–86 (2005).
92. Wright DE, Wang C-Y, Kao C-F. Histone ubiquitylation and chromatin dynamics. Front. Biosci. (Landmark Ed. 17, 1051–78 (2012).
93. Osley MA. Regulation of histone H2A and H2B ubiquitylation. Brief. Funct. Genomic. Proteomic. 5(3), 179–89 (2006).
94. Weake VM, Workman JL. Histone ubiquitination: triggering gene activity. Mol. Cell. 29(6), 653–63 (2008).
95. Nathan D, Sterner DE, Berger SL. Histone modifications: Now summoning sumoylation. Proc. Natl. Acad. Sci. 100(23), 13118–13120 (2003).
96. Iñiguez-Lluhí JA. For a Healthy Histone Code, a Little SUMO in the Tail Keeps the Acetyl Away. ACS Chem. Biol. 1(4), 204–206 (2006).
97. Shiio Y, Eisenman RN. Histone sumoylation is associated with transcriptional repression. Proc. Natl. Acad. Sci. U. S. A. 100(23), 13225–30 (2003).
98. Gill G. Something about SUMO inhibits transcription. Curr. Opin. Genet. Dev. 15(5), 536–41 (2005).
99. Haery L, Thompson RC, Gilmore TD. Histone acetyltransferases and histone deacetylases in B- and T-cell development, physiology and malignancy. Genes Cancer. 6(5-6), 184–213 (2015).
100. Zhang C, Zhong JF, Stucky A, Chen X-L, Press MF, Zhang X. Histone acetylation: novel target for the treatment of acute lymphoblastic leukemia. Clin. Epigenetics. 7(1), 117 (2015).
101. Josling GA, Selvarajah SA, Petter M, Duffy MF. The role of bromodomain proteins in regulating gene expression. Genes (Basel). 3(2), 320–43 (2012).
181
102. Dokmanovic M, Clarke C, Marks PA. Histone deacetylase inhibitors: overview and perspectives. Mol. Cancer Res. 5(10), 981–9 (2007).
103. Zhang T, Cooper S, Brockdorff N. The interplay of histone modifications - writers that read. EMBO Rep. 16(11), 1467–81 (2015).
104. Liang G, Lin JCY, Wei V, et al. Distinct localization of histone H3 acetylation and H3-K4 methylation to the transcription start sites in the human genome. Proc. Natl. Acad. Sci. U. S. A. 101(19), 7357–62 (2004).
105. Shaknovich R, Melnick A. Epigenetics and B-cell lymphoma. Curr. Opin. Hematol. 18(4), 293–9 (2011).
106. Herz H-M, Garruss A, Shilatifard A. SET for life: biochemical activities and biological functions of SET domain-containing proteins. Trends Biochem. Sci. 38(12), 621–39 (2013).
107. Moore KE, Gozani O. An unexpected journey: lysine methylation across the proteome. Biochim. Biophys. Acta. 1839(12), 1395–403 (2014).
108. Rivera C, Gurard-Levin ZA, Almouzni G, Loyola A. Histone lysine methylation and chromatin replication. Biochim. Biophys. Acta - Gene Regul. Mech. 1839(12), 1433– 1439 (2014).
109. Yun M, Wu J, Workman JL, Li B. Readers of histone modifications. Cell Res. 21(4), 564–578 (2011).
110. Wolf SS. The protein arginine methyltransferase family: an update about function, new perspectives and the physiological role in humans. Cell Mol Life Sci. 66(13), 2109–2121 (2009).
111. Yang Y, Hadjikyriacou A, Xia Z, et al. PRMT9 is a type II methyltransferase that methylates the splicing factor SAP145. Nat. Commun. 6, 6428 (2015).
112. Zurita-Lopez CI, Sandberg T, Kelly R, Clarke SG. Human protein arginine 182
methyltransferase 7 (PRMT7) is a type III enzyme forming ω-NG-monomethylated arginine residues. J. Biol. Chem. 287(11), 7859–70 (2012).
113. Migliori V, Müller J, Phalke S, et al. Symmetric dimethylation of H3R2 is a newly identified histone mark that supports euchromatin maintenance. Nat. Struct. Mol. Biol. 19(2), 136–44 (2012).
114. Shi YG, Tsukada Y. The discovery of histone demethylases. Cold Spring Harb. Perspect. Biol. 5(9) (2013).
115. McGrath J, Trojer P. Targeting histone lysine methylation in cancer. Pharmacol. Ther. 150, 1–22 (2015).
116. Liu K, Liu Y, Lau JL, Min J. Epigenetic targets and drug discovery Part 2: Histone demethylation and DNA methylation. Pharmacol. Ther. 151, 121–40 (2015).
117. Li KK, Luo C, Wang D, Jiang H, Zheng YG. Chemical and biochemical approaches in the study of histone methylation and demethylation. Med. Res. Rev. 32(4), 815–67 (2012).
118. Chang B, Chen Y, Zhao Y, Bruick RK. JMJD6 is a histone arginine demethylase. Science. 318(5849), 444–7 (2007).
119. Muntean AG, Hess JL, Holliday R, et al. Epigenetic dysregulation in cancer. Am. J. Pathol. 175(4), 1353–61 (2009).
120. Zhao S, Geybels MS, Leonardson A, et al. Epigenome-wide Tumor DNA Methylation Profiling Identifies Novel Prognostic Biomarkers of Metastatic-lethal Progression in Men with Clinically Localized Prostate Cancer. Clin. Cancer Res. , clincanres.0549.2016 (2016).
121. O’Sullivan DE, Johnson KC, Skinner L, Koestler DC, Christensen BC. Epigenetic and genetic burden measures are associated with tumor characteristics in invasive breast carcinoma. http://dx.doi.org/10.1080/15592294.2016.1168673. (2016).
183
122. Kong W, He L, Richards EJ, et al. Upregulation of miRNA-155 promotes tumour angiogenesis by targeting VHL and is associated with poor prognosis and triple- negative breast cancer. Oncogene. 33(6), 679–689 (2014).
123. Zeng Q, Tao X, Huang F, et al. Overexpression of miR-155 promotes the proliferation and invasion of oral squamous carcinoma cells by regulating BCL6/cyclin D2. Int. J. Mol. Med. 37(5), 1274–80 (2016).
124. Bhalla KN, Fiskus W. NEDD8 and HDACs: promising cotargets in AML. Blood. 127(18), 2168–70 (2016).
125. Pal S, Vishwanath SN, Erdjument-Bromage H, Tempst P, Sif S. Human SWI/SNF- associated PRMT5 methylates histone H3 arginine 8 and negatively regulates expression of ST7 and NM23 tumor suppressor genes. Mol. Cell. Biol. 24(21), 9630–45 (2004).
126. Zinzalla G. A New Way Forward in Cancer Drug Discovery: Inhibiting the SWI/SNF Chromatin Remodelling Complex. ChemBioChem. 17(8), 677–682 (2016).
127. Pal S, Baiocchi RA, Byrd JC, Grever MR, Jacob ST, Sif S. Low levels of miR-92b/96 induce PRMT5 translation and H3R8/H4R3 methylation in mantle cell lymphoma. EMBO J. 26(15), 3558–69 (2007).
128. Antonysamy S, Bonday Z, Campbell RM, et al. Crystal structure of the human PRMT5:MEP50 complex. Proc. Natl. Acad. Sci. U. S. A. 109(44), 17960–5 (2012).
129. Wang M, Xu R-M, Thompson PR. Substrate specificity, processivity, and kinetic mechanism of protein arginine methyltransferase 5. Biochemistry. 52(32), 5430–40 (2013).
130. Pal S, Vishwanath SN, Erdjument-Bromage H, Tempst P, Sif S. Human SWI/SNF- associated PRMT5 methylates histone H3 arginine 8 and negatively regulates expression of ST7 and NM23 tumor suppressor genes. Mol Cell Biol. 24(21), 9630– 9645 (2004).
131. Ancelin K, Lange UC, Hajkova P, et al. Blimp1 associates with Prmt5 and directs 184
histone arginine methylation in mouse germ cells. Nat. Cell Biol. 8(6), 623–30 (2006).
132. Stopa N, Krebs JE, Shechter D. The PRMT5 arginine methyltransferase: many roles in development, cancer and beyond. Cell. Mol. Life Sci. (2015).
133. Tee W-W, Pardo M, Theunissen TW, et al. Prmt5 is essential for early mouse development and acts in the cytoplasm to maintain ES cell pluripotency. Genes Dev. 24(24), 2772–7 (2010).
134. Bezzi M, Teo SX, Muller J, et al. Regulation of constitutive and alternative splicing by PRMT5 reveals a role for Mdm4 pre-mRNA in sensing defects in the spliceosomal machinery. Genes Dev. 27(17), 1903–16 (2013).
135. Wang Y, Zhu T, Li Q, et al. Prmt5 is required for germ cell survival during spermatogenesis in mice. Sci. Rep. 5, 11031 (2015).
136. Wang Y, Li Q, Liu C, et al. Protein Arginine Methyltransferase 5 (Prmt5) Is Required for Germ Cell Survival During Mouse Embryonic Development. Biol. Reprod. 92(4), 104–104 (2015).
137. Kim S, Günesdogan U, Zylicz JJ, et al. PRMT5 protects genomic integrity during global DNA demethylation in primordial germ cells and preimplantation embryos. Mol. Cell. 56(4), 564–79 (2014).
138. Lee YH, Stallcup MR. Minireview: protein arginine methylation of nonhistone proteins in transcriptional regulation. Mol Endocrinol. 23(4), 425–433 (2009).
139. Bandyopadhyay S, Harris DP, Adams GN, et al. HOXA9 methylation by PRMT5 is essential for endothelial cell expression of leukocyte adhesion molecules. Mol. Cell. Biol. 32(7), 1202–13 (2012).
140. Wei H, Wang B, Miyagi M, et al. PRMT5 dimethylates R30 of the p65 subunit to activate NF-κB. Proc. Natl. Acad. Sci. U. S. A. 110(33), 13516–21 (2013).
185
141. Harris DP, Bandyopadhyay S, Maxwell TJ, Willard B, Dicorleto PE. TNF-α Induction of CXCL10 in Endothelial Cells Requires Protein Arginine Methyltransferase 5 (PRMT5)-mediated NF-κB p65 Methylation. J. Biol. Chem. 289(22), 15328–15339 (2014).
142. Jansson M, Durant ST, Cho EC, et al. Arginine methylation regulates the p53 response. Nat Cell Biol. 10(12), 1431–1439 (2008).
143. Scoumanne A, Zhang J, Chen X. PRMT5 is required for cell-cycle progression and p53 tumor suppressor function. Nucleic Acids Res. 37(15), 4965–4976 (2009).
144. Zhang T, Günther S, Looso M, et al. Prmt5 is a regulator of muscle stem cell expansion in adult mice. Nat. Commun. 6, 7140 (2015).
145. Beltran-Alvarez P, Espejo A, Schmauder R, et al. Protein arginine methyl transferases-3 and -5 increase cell surface expression of cardiac sodium channel. FEBS Lett. 587(19), 3159–65 (2013).
146. Liu F, Cheng G, Hamard P-J, et al. Arginine methyltransferase PRMT5 is essential for sustaining normal adult hematopoiesis. J. Clin. Invest. 125(125(9)) (2015).
147. Richard S, Morel M, Cléroux P. Arginine methylation regulates IL-2 gene expression: a role for protein arginine methyltransferase 5 (PRMT5). Biochem. J. 388(Pt 1), 379–386 (2005).
148. Paul C, Sardet C, Fabbrizio E. The Wnt-target gene Dlk-1 is regulated by the Prmt5-associated factor Copr5 during adipogenic conversion. Biol. Open. 4(3), 312–316 (2015).
149. LeBlanc SE, Konda S, Wu Q, et al. Protein arginine methyltransferase 5 (Prmt5) promotes gene expression of peroxisome proliferator-activated receptor γ2 (PPARγ2) and its target genes during adipogenesis. Mol. Endocrinol. 26(4), 583–97 (2012).
150. Zhou Z, Sun X, Zou Z, et al. PRMT5 regulates Golgi apparatus structure through methylation of the golgin GM130. Cell Res. 20(9), 1023–1033 (2010). 186
151. Saha K, Eckert RL. Methylosome Protein 50 and PKCδ/p38δ Protein Signaling Control Keratinocyte Proliferation via Opposing Effects on p21Cip1 Gene Expression. J. Biol. Chem. 290(21), 13521–30 (2015).
152. Huang J, Vogel G, Yu Z, Almazan G, Richard S. Type II arginine methyltransferase PRMT5 regulates gene expression of inhibitors of differentiation/DNA binding Id2 and Id4 during glial cell differentiation. J. Biol. Chem. 286(52), 44424–32 (2011).
153. Wei T-YW, Juan C-C, Hisa J-Y, et al. Protein arginine methyltransferase 5 is a potential oncoprotein that upregulates G1 cyclins/cyclin-dependent kinases and the phosphoinositide 3-kinase/AKT signaling cascade. Cancer Sci. 103(9), 1640–50 (2012).
154. Tarighat SS, Santhanam R, Frankhouser D, et al. The dual epigenetic role of PRMT5 in acute myeloid leukemia: gene activation and repression via histone arginine methylation. Leukemia. (2015).
155. Chung J, Karkhanis V, Tae S, et al. Protein arginine methyltransferase 5 (PRMT5) inhibition induces lymphoma cell death through reactivation of the retinoblastoma tumor suppressor pathway and polycomb repressor complex 2 (PRC2) silencing. J. Biol. Chem. 288(49), 35534–47 (2013).
156. American Cancer Society: Cancer Facts and Figures 2016. [Internet]. Atlanta, GA Am. Cancer Soc. (2016). Available from: http://www.cancer.gov/types/common- cancers.
157. Han X, Li R, Zhang W, et al. Expression of PRMT5 correlates with malignant grade in gliomas and plays a pivotal role in tumor growth in vitro. J. Neurooncol. 118(1), 61–72 (2014).
158. Yan F, Alinari L, Lustberg ME, et al. Genetic validation of the protein arginine methyltransferase PRMT5 as a candidate therapeutic target in glioblastoma. Cancer Res. 74(6), 1752–65 (2014).
159. What is nasopharyngeal cancer? [Internet]. Atlanta, GA Am. Cancer Soc. (2015). Available from:
187
http://www.cancer.org/cancer/nasopharyngealcancer/detailedguide/nasopharyng eal-cancer-what-is-nasopharyngeal-cancer.
160. Yang D, Liang T, Gu Y, et al. Protein N-arginine methyltransferase 5 promotes the tumor progression and radioresistance of nasopharyngeal carcinoma. Oncol. Rep. 35(3), 1703–1710 (2016).
161. Shilo K, Wu X, Sharma S, et al. Cellular localization of protein arginine methyltransferase-5 correlates with grade of lung tumors. Diagn. Pathol. 8(1), 201 (2013).
162. Gu Z, Gao S, Zhang F, et al. Protein arginine methyltransferase 5 is essential for growth of lung cancer cells. Biochem. J. 446(2), 235–41 (2012).
163. Ibrahim R, Matsubara D, Osman W, et al. Expression of PRMT5 in lung adenocarcinoma and its significance in epithelial-mesenchymal transition. Hum. Pathol. 45(7), 1397–405 (2014).
164. Yang F, Wang J, Ren H-Y, et al. Proliferative role of TRAF4 in breast cancer by upregulating PRMT5 nuclear expression. Tumour Biol. 36(8), 5901–11 (2015).
165. Powers MA, Fay MM, Factor RE, Welm AL, Ullman KS. Protein arginine methyltransferase 5 accelerates tumor growth by arginine methylation of the tumor suppressor programmed cell death 4. Cancer Res. 71(16), 5579–87 (2011).
166. Kohler BA, Sherman RL, Howlader N, et al. Annual Report to the Nation on the Status of Cancer, 1975-2011, Featuring Incidence of Breast Cancer Subtypes by Race/Ethnicity, Poverty, and State. J. Natl. Cancer Inst. 107(6), djv048 (2015).
167. Bao X, Zhao S, Liu T, Liu Y, Liu Y, Yang X. Overexpression of PRMT5 promotes tumor cell growth and is associated with poor disease prognosis in epithelial ovarian cancer. J. Histochem. Cytochem. 61(3), 206–17 (2013).
168. Gu Z, Li Y, Lee P, Liu T, Wan C, Wang Z. Protein arginine methyltransferase 5 functions in opposite ways in the cytoplasm and nucleus of prostate cancer cells. PLoS One. 7(8), e44033 (2012). 188
169. Zhang B, Dong S, Li Z, et al. Targeting protein arginine methyltransferase 5 inhibits human hepatocellular carcinoma growth via the downregulation of beta-catenin. J. Transl. Med. 13(1), 349 (2015).
170. Liu L, Zhao X, Zhao L, et al. Arginine Methylation of SREBP1a via PRMT5 Promotes De Novo Lipogenesis and Tumor Growth. Cancer Res. 76(5), 1260–72 (2016).
171. Zhang B, Dong S, Zhu R, et al. Targeting protein arginine methyltransferase 5 inhibits colorectal cancer growth by decreasing arginine methylation of eIF4E and FGFR3. Oncotarget. 5 (2015).
172. Pak MG, Lee HW, Roh MS. High nuclear expression of protein arginine methyltransferase-5 is a potentially useful marker to estimate submucosal invasion in endoscopically resected early colorectal carcinoma. Pathol. Int. 65(10), 541–8 (2015).
173. Nicholas C, Yang J, Peters SB, et al. PRMT5 is upregulated in malignant and metastatic melanoma and regulates expression of MITF and p27(Kip1.). PLoS One. 8(9), e74710 (2013).
174. Park JH, Szemes M, Vieira GC, et al. Protein arginine methyltransferase 5 is a key regulator of the MYCN oncoprotein in neuroblastoma cells. Mol. Oncol. 9(3), 617– 627 (2014).
175. Li Y, Chitnis N, Nakagawa H, et al. PRMT5 Is Required for Lymphomagenesis Triggered by Multiple Oncogenic Drivers. Cancer Discov. 5(3), 288–303 (2015).
176. Aggarwal P, Vaites LP, Kim JK, et al. Nuclear cyclin D1/CDK4 kinase regulates CUL4 expression and triggers neoplastic growth via activation of the PRMT5 methyltransferase. Cancer Cell. 18(4), 329–340 (2010).
177. Panfil AR, Al-Saleem J, Howard CM, et al. PRMT5 Is Upregulated in HTLV-1- Mediated T-Cell Transformation and Selective Inhibition Alters Viral Gene Expression and Infected Cell Survival. Viruses. 8(1), 7 (2015).
178. Ho M-C, Wilczek C, Bonanno JB, et al. Structure of the arginine methyltransferase 189
PRMT5-MEP50 reveals a mechanism for substrate specificity. PLoS One. 8(2), e57008 (2013).
179. Lacroix M, El Messaoudi S, Rodier G, Le Cam A, Sardet C, Fabbrizio E. The histone- binding protein COPR5 is required for nuclear functions of the protein arginine methyltransferase PRMT5. EMBO Rep. 9(5), 452–8 (2008).
180. Lacroix M, El Messaoudi S, Rodier G, Le Cam A, Sardet C, Fabbrizio E. The histone- binding protein COPR5 is required for nuclear functions of the protein arginine methyltransferase PRMT5. EMBO Rep. 9(5), 452–458 (2008).
181. Pesiridis GS, Diamond E, Van Duyne GD. Role of pICLn in methylation of Sm proteins by PRMT5. J. Biol. Chem. 284(32), 21347–59 (2009).
182. Pellizzoni L, Yong J, Dreyfuss G. Essential role for the SMN complex in the specificity of snRNP assembly. Science. 298(5599), 1775–9 (2002).
183. Yanling Zhao D, Gish G, Braunschweig U, et al. SMN and symmetric arginine dimethylation of RNA polymerase II C-terminal domain control termination. Nature. 529(7584), 48–53 (2015).
184. Guderian G, Peter C, Wiesner J, et al. RioK1, a new interactor of protein arginine methyltransferase 5 (PRMT5), competes with pICln for binding and modulates PRMT5 complex composition and substrate specificity. J. Biol. Chem. 286(3), 1976– 86 (2011).
185. Gurung B, Feng Z, Iwamoto D V, et al. Menin epigenetically represses Hedgehog signaling in MEN1 tumor syndrome. Cancer Res. 73(8), 2650–8 (2013).
186. Gurung B, Hua X. Menin/PRMT5/hedgehog signaling: a potential target for the treatment of multiple endocrine neoplasia type 1 tumors. Epigenomics. 5(5), 469– 71 (2013).
187. Kirino Y, Kim N, de Planell-Saguer M, et al. Arginine methylation of Piwi proteins catalysed by dPRMT5 is required for Ago3 and Aub stability. Nat. Cell Biol. 11(5), 652–8 (2009). 190
188. Vagin V V, Wohlschlegel J, Qu J, et al. Proteomic analysis of murine Piwi proteins reveals a role for arginine methylation in specifying interaction with Tudor family members. Genes Dev. 23(15), 1749–62 (2009).
189. Hou Z, Peng H, Ayyanathan K, et al. The LIM protein AJUBA recruits protein arginine methyltransferase 5 to mediate SNAIL-dependent transcriptional repression. Mol. Cell. Biol. 28(10), 3198–207 (2008).
190. Pal S, Yun R, Datta A, et al. mSin3A/histone deacetylase 2- and PRMT5-containing Brg1 complex is involved in transcriptional repression of the Myc target gene cad. Mol Cell Biol. 23(21), 7475–7487 (2003).
191. Seth-Vollenweider T, Joshi S, Dhawan P, Sif S, Christakos S. Novel mechanism of negative regulation of 1,25-dihydroxyvitamin D3-induced 25-hydroxyvitamin D3 24-hydroxylase (Cyp24a1) Transcription: epigenetic modification involving cross- talk between protein-arginine methyltransferase 5 and the SWI/SNF complex. J. Biol. Chem. 289(49), 33958–70 (2014).
192. Wang Y, Wysocka J, Sayegh J, et al. Human PAD4 regulates histone arginine methylation levels via demethylimination. Science. 306(5694), 279–83 (2004).
193. Cuthbert GL, Daujat S, Snowden AW, et al. Histone deimination antagonizes arginine methylation. Cell. 118(5), 545–53 (2004).
194. Pal S, Baiocchi RA, Byrd JC, Grever MR, Jacob ST, Sif S. Low levels of miR-92b/96 induce PRMT5 translation and H3R8/H4R3 methylation in mantle cell lymphoma. EMBO J. 26(15), 3558–3569 (2007).
195. Zhang H-T, Zhang D, Zha Z-G, Hu C-D. Transcriptional activation of PRMT5 by NF- Y is required for cell growth and negatively regulated by the PKC/c-Fos signaling in prostate cancer cells. Biochim. Biophys. Acta. 1839(11), 1330–40 (2014).
196. Yang Y, Bedford MT. Protein arginine methyltransferases and cancer. Nat. Rev. Cancer. 13(1), 37–50 (2013).
197. Zhang H-T, Zeng L-F, He Q-Y, Tao WA, Zha Z-G, Hu C-D. The E3 ubiquitin ligase 191
CHIP mediates ubiquitination and proteasomal degradation of PRMT5. Biochim. Biophys. Acta. 1863(2), 335–346 (2015).
198. Smil D, Eram MS, Li F, et al. Discovery of a Dual PRMT5–PRMT7 Inhibitor. ACS Med. Chem. Lett. 6(4), 150305065104002 (2015).
199. Alinari L, Mahasenan K V, Yan F, et al. Selective inhibition of protein arginine methyltransferase 5 blocks initiation and maintenance of B-cell transformation. Blood. 125(16), 2530–2543 (2015).
200. Chan-Penebre E, Kuplast KG, Majer CR, et al. A selective inhibitor of PRMT5 with in vivo and in vitro potency in MCL models. Nat. Chem. Biol. 11(6), 432–437 (2015).
201. Hu H, Owens EA, Su H, et al. Exploration of cyanine compounds as selective inhibitors of protein arginine methyltransferases: synthesis and biological evaluation. J. Med. Chem. 58(3), 1228–43 (2015).
202. Jares P, Colomer D, Campo E. Genetic and molecular pathogenesis of mantle cell lymphoma: perspectives for new targeted therapeutics. Nat Rev Cancer. 7(10), 750–762 (2007).
203. Shah BD, Martin P, Sotomayor EM. Mantle cell lymphoma: a clinically heterogeneous disease in need of tailored approaches. Cancer Control. 19(3), 227– 35 (2012).
204. Salaverria I, Perez-galan P, Colomer D, Campo E. Mantle cell lymphoma: from pathology and molecular pathogenesis to new therapeutic perspectives. Hematol. J. 90(10), 11–16 (2005).
205. Pérez-Galán P, Dreyling M, Wiestner A. Mantle cell lymphoma: biology, pathogenesis, and the molecular basis of treatment in the genomic era. Blood. 117(1), 26–38 (2011).
206. Vose JM. Mantle cell lymphoma: 2015 update on diagnosis, risk-stratification, and clinical management. Am. J. Hematol. 90(8), 739–45 (2015).
192
207. Dreyling M. Mantle cell lymphoma: biology, clinical presentation, and therapeutic approaches. Am. Soc. Clin. Oncol. Educ. Book. , 191–8 (2014).
208. Skarbnik AP, Smith MR. Therapies for Mantle Cell Lymphoma: Current Challenges and a Brighter Future. Discov. Med. 15(82), 177–187 (2013).
209. Dreyling M, Kluin-Nelemans HC, Beà S, et al. Update on the molecular pathogenesis and clinical treatment of mantle cell lymphoma: report of the 11th annual conference of the European Mantle Cell Lymphoma Network. Leuk. Lymphoma. 54(4), 699–707 (2013).
210. Bodrug SE, Warner BJ, Bath ML, Lindeman GJ, Harris AW, Adams JM. Cyclin D1 transgene impedes lymphocyte maturation and collaborates in lymphomagenesis with the myc gene. EMBO J. 13(9), 2124–2130 (1994).
211. Jares P, Colomer D, Campo E. Molecular pathogenesis of mantle cell lymphoma. J. Clin. Invest. 122(10), 3416–23 (2012).
212. Bea S, Tort F, Pinyol M, et al. BMI-1 Gene Amplification and Overexpression in Hematological Malignancies Occur Mainly in Mantle Cell Lymphomas. Cancer Res. 61(6), 2409–2412 (2001).
213. Weniger MA, Wiestner A. Molecular targeted approaches in mantle cell lymphoma. Semin Hematol. 48(3), 214–226 (2011).
214. Martin P, Chadburn A, Christos P, et al. Outcome of deferred initial therapy in mantle-cell lymphoma. J. Clin. Oncol. 27(8), 1209–13 (2009).
215. Dreyling M, Thieblemont C, Gallamini A, et al. ESMO Consensus conferences: guidelines on malignant lymphoma. part 2: marginal zone lymphoma, mantle cell lymphoma, peripheral T-cell lymphoma. Ann. Oncol. 24(4), 857–77 (2013).
216. Dreyling M, Lenz G, Hoster E, et al. Early consolidation by myeloablative radiochemotherapy followed by autologous stem cell transplantation in first remission significantly prolongs progression-free survival in mantle-cell lymphoma: results of a prospective randomized trial of the European . Blood. 193
105(7), 2677–84 (2005).
217. Humala K, Younes A. Current and emerging new treatment strategies for mantle cell lymphoma. Leuk. Lymphoma. 54(5), 912–21 (2013).
218. Romaguera JE, Fayad LE, Feng L, et al. Ten-year follow-up after intense chemoimmunotherapy with Rituximab-HyperCVAD alternating with Rituximab- high dose methotrexate/cytarabine (R-MA) and without stem cell transplantation in patients with untreated aggressive mantle cell lymphoma. Br. J. Haematol. 150(2), 200–8 (2010).
219. Cheah CY, Seymour JF, Wang ML. Mantle Cell Lymphoma. J. Clin. Oncol. , JCO.2015.63.5904– (2016).
220. Wang L, Pal S, Sif S. Protein arginine methyltransferase 5 suppresses the transcription of the RB family of tumor suppressors in leukemia and lymphoma cells. Mol Cell Biol. 28(20), 6262–6277 (2008).
221. Avivi I, Goy A. Refining the Mantle Cell Lymphoma Paradigm: Impact of Novel Therapies on Current Practice. Clin. Cancer Res. 21(17), 3853–61 (2015).
222. Liang JJ, Wang Z, Chiriboga L, et al. The expression and function of androgen receptor coactivator p44 and protein arginine methyltransferase 5 in the developing testis and testicular tumors. J. Urol. 177(5), 1918–22 (2007).
223. Rudelius M, Pittaluga S, Nishizuka S, et al. Constitutive activation of Akt contributes to the pathogenesis and survival of mantle cell lymphoma. Blood. 108(5), 1668–76 (2006).
224. Andreu-Pérez P, Esteve-Puig R, de Torre-Minguela C, et al. Protein arginine methyltransferase 5 regulates ERK1/2 signal transduction amplitude and cell fate through CRAF. Sci. Signal. 4(190), ra58 (2011).
225. Dennison JB, Shanmugam M, Ayres ML, et al. 8-Aminoadenosine inhibits Akt/mTOR and Erk signaling in mantle cell lymphoma. Blood. 116(25), 5622–30 (2010). 194
226. Fu K, Weisenburger DD, Greiner TC, et al. Cyclin D1-negative mantle cell lymphoma: a clinicopathologic study based on gene expression profiling. Blood. 106(13), 4315–21 (2005).
227. Alinari L, White VL, Earl CT, et al. Combination bortezomib and rituximab treatment affects multiple survival and death pathways to promote apoptosis in mantle cell lymphoma. MAbs. 1(1), 31–40 (2009).
228. Setoodeh R, Schwartz S, Papenhausen P, et al. Double-hit mantle cell lymphoma with MYC gene rearrangement or amplification: a report of four cases and review of the literature. Int. J. Clin. Exp. Pathol. 6(2), 155–67 (2013).
229. Rouaud P, Fiancette R, Vincent-Fabert C, et al. Mantle cell lymphoma-like lymphomas in c-myc-3’RR/p53+/- mice and c-myc-3'RR/Cdk4R24C mice: differential oncogenic mechanisms but similar cellular origin. Oncotarget. 3(5), 586–93 (2012).
230. Zhang X-F, Li K, Gao L, et al. miR-191 promotes tumorigenesis of human colorectal cancer through targeting C/EBPβ. Oncotarget. 6(6), 4144–4158 (2015).
231. Tsutsui T, Fukasawa R, Shinmyouzu K, et al. Mediator complex recruits epigenetic regulators via its two cyclin-dependent kinase subunits to repress transcription of immune response genes. J. Biol. Chem. 288(29), 20955–65 (2013).
232. Zhen Y, Liu Z, Yang H, et al. Tumor suppressor PDCD4 modulates miR-184- mediated direct suppression of C-MYC and BCL2 blocking cell growth and survival in nasopharyngeal carcinoma. Cell Death Dis. 4, e872 (2013).
233. Zhao Q, Rank G, Tan YT, et al. PRMT5-mediated methylation of histone H4R3 recruits DNMT3A, coupling histone and DNA methylation in gene silencing. Nat Struct Mol Biol. 16(3), 304–311 (2009).
234. Jansson M, Durant ST, Cho E-C, et al. Arginine methylation regulates the p53 response. Nat. Cell Biol. 10(12), 1431–9 (2008).
235. Guo Y, Stacey DW, Hitomi M. Post-transcriptional regulation of cyclin D1 195
expression during G2 phase. Oncogene. 21(49), 7545–7556 (2002).
236. Lim J-H, Lee Y-M, Lee G, et al. PRMT5 is essential for the eIF4E-mediated 5’-cap dependent translation. Biochem. Biophys. Res. Commun. 452(4), 1016–21 (2014).
237. Koh CM, Bezzi M, Low DHP, et al. MYC regulates the core pre-mRNA splicing machinery as an essential step in lymphomagenesis. Nature. 523(7558), 96–100 (2015).
238. Wilkie TM, Brinster RL, Palmiter RD. Germline and somatic mosaicism in transgenic mice. Dev. Biol. 118(1), 9–18 (1986).
239. Harris AW, Pinkert CA, Crawford M, Langdon WY, Brinster RL, Adams JM. The E mu-myc transgenic mouse. A model for high-incidence spontaneous lymphoma and leukemia of early B cells. J. Exp. Med. 167(2), 353–71 (1988).
240. Gladden AB, Woolery R, Aggarwal P, Wasik MA, Diehl JA. Expression of constitutively nuclear cyclin D1 in murine lymphocytes induces B-cell lymphoma. Oncogene. 25(7), 998–1007 (2006).
241. Grosschedl R, Weaver D, Baltimore D, et al. Introduction of a μ immunoglobulin gene into the mouse germ line: Specific expression in lymphoid cells and synthesis of functional antibody. Cell. 38(3), 647–658 (1984).
242. Mowen KA, Schurter BT, Fathman JW, David M, Glimcher LH. Arginine Methylation of NIP45 Modulates Cytokine Gene Expression in Effector T Lymphocytes. Mol. Cell. 15(4), 559–571 (2004).
243. Lymphomainfo.net. http://www.lymphomainfo.net/articles/non-hodgkins/t-cell/t- cell-lymphoma-prognosis [Internet]. (2013). Available from: http://www.lymphomainfo.net/articles/non-hodgkins/t-cell/t-cell-lymphoma- prognosis.
244. Li Y, Diehl JA. PRMT5-dependent p53 escape in tumorigenesis. Oncoscience. 2(8), 700–2 (2015).
196
245. Tae S, Karkhanis V, Velasco K, et al. Bromodomain protein 7 interacts with PRMT5 and PRC2, and is involved in transcriptional repression of their target genes. Nucleic Acids Res. 39(13), 5424–5438 (2011).
246. Bose RJC, Lee S-H, Park H. Biofunctionalized nanoparticles: an emerging drug delivery platform for various disease treatments. Drug Discov. Today. (2016).
247. Jin CY, Han MH, Lee SS, Choi YH. Mass producible and biocompatible microneedle patch and functional verification of its usefulness for transdermal drug delivery. Biomed. Microdevices. 11(6), 1195–1203 (2009).
248. Chen M, Yi B, Sun J. Inhibition of cardiomyocyte hypertrophy by protein arginine methyltransferase 5. J. Biol. Chem. 289(35), 24325–35 (2014).
249. Bodrug SE, Warner BJ, Bath ML, Lindeman GJ, Harris AW, Adams JM. Cyclin D1 transgene impedes lymphocyte maturation and collaborates in lymphomagenesis with the myc gene. EMBO J. 13(9), 2124–30 (1994).
250. Ondrejka SL, Hsi ED. T-cell Lymphomas: Updates in Biology and Diagnosis. Surg. Pathol. Clin. 9(1), 131–141 (2016).
251. Mahler JF, Stokes W, Mann PC, Takaoka M, Maronpot RR. Spontaneous lesions in aging FVB/N mice. Toxicol. Pathol. 24(6), 710–6.
197
Appendix A: List of Abbreviations
5mc 5-methylcytosine
ADMA asymmetric dimethyl arginine
BLIMP1 B lymphocyte induced maturation protein 1
BRD bromodomain
CD cluster of differentiation
CDK cyclin-dependent kinase
CDKN cyclin-dependent kinase inhibitor
CEBPβ CCAAT/enhancer binding protein beta
ChIP chromatin immunoprecipitation
Co-IP co-immunoprecipitation
COPR5 cooperator of PRMT5
DNMT DNA methyltransferase
DUBs deubiquitinating enzymes
ECOG Eastern Cooperative Oncology Group
EMT epithelial-mesenchymal transition
FACS fluorescence activated cell sorting
HATs histone acetyltransferases
198
HDAC histone deacetylase
HMTs histone methyltransferases hPRMT5 human PRMT5
Ig immunoglobulin
IL interleukin
LDH lactic dehydrogenase
MCL mantle cell lymphoma
MEP50 methylosome protein 50
MIDI Mantle cell International Prognostic Index
NATs normal adjacent tissues
NHL non-Hodgkin’s lymphoma nTg non-transgenic
OS overall survival
PFS progression free survival
PRMT protein arginine methyltransferase
PRMT5 protein arginine methyltransferase 5
RB retinoblastoma
SAM s-adenosyl methionine
SDMA symmetric dimethyl arginine
ST7 suppressor of tumorigenicity 7
TCR T-cell receptor
199
Tg transgenic
UTR untranslated region
WT wild type
200