Systematic Functional Proteomic Investigation of Mammalian Histone Methylation-Related Protein Complexes

Jonathan B. Olsen

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Graduate Department of Molecular Genetics University of Toronto

Investigation of Histone Methylation-Related Protein Complexes

Jonathan B. Olsen

Doctor of Philosophy

Molecular Genetics University of Toronto

2014 Abstract

Epigenetic regulation of chromatin structure, which involves site-specific methylation of histone proteins, is critical for regulating gene expression during development. While the enzymatic components of histone methylation-related protein complexes have been discovered, little is generally known regarding the associated tissue-specific co-factors that regulate complex activity in vertebrate/mammalian embryos. To better understand the composition and control of histone methylation-related complexes in mammals, I used an affinity purification and mass spectrometry (AP-MS) approach to identity stably interacting proteins of histone methylation- related proteins in human embryonic kidney (HEK) 293 cells. My AP-MS approach included replicate isolation and characterization of 33 histone methylation- and/or transcription-related proteins, resulting in an interaction network encompassing 573 protein-protein interactions

(PPIs). Among the findings were a novel set of RNA polymerase II (RNAPII) binding proteins, known as the “RPRD” proteins, which recognize phosphorylated RNAPII, and several putative new components of the evolutionarily conserved gene silencing machinery. Most notably, I identified and characterized ZNF644, a protein that is putatively causally linked to high-grade myopia, as a novel interacting partner for the paralogous histone methyltransferases (HMTs) G9a and GLP and as a new co-regulator of histone H3 at lysine 9 dimethylation (H3K9me2). Using a zebrafish embryo model system, I characterized the roles of zebrafish g9a and two orthologues of ii

ZNF644, termed znf644a and znf644b, in the forming retina and midbrain. I found that even modest disruption of g9a activity caused widespread apoptosis in progenitor and differentiating cell populations in the retina. I also found that znf644a and znf644b have common retina-specific roles in suppressing progenitor gene expression (i.e., vsx2 and ccnd1) by regulating promoter

H3K9me2. However, despite the overlapping roles in progenitor gene suppression, the two znf644 paralogues controlled distinct aspects of retinal differentiation; specifically, I observed that disruption of znf644b caused retina-specific developmental delays, resulting in the delayed differentiation of RPCs, whereas disruption of znf644a caused the mislocalized expression of

H3K9me2 marked genes (i.e., znf644a and znf644b) during differentiation, resulting in the impaired viability of differentiated neurons. Importantly, I used complementation assays as well as “genetic cooperativity” assays to demonstrate that, despite the disparate phenotypes of g9a, znf644a and znf644b morphant retinas, the regulatory roles of znf644a and znf644b are critically dependent on their ability to functionally and physically interact with g9a. In addition, I provided evidence that the functions of znf644a and znf644b are encompassed by mammalian ZNF644 and that the retinal roles of g9a, znf644a and znf644b are recapitulated in progenitor cells of the developing midbrain. Collectively, my thesis provides both a high-confidence global molecular interaction map that furthers understanding of the physical and functional associations underlying the mammalian histone methylation machinery and highlights the unexpected role of a previously unappreciated co-factor, ZNF644 as a co-regulator of H3K9me2-mediated gene silencing and a multifaceted regulator of neural differentiation.

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Acknowledgments

The work in this doctoral dissertation was supported in part by grants from the Canadian

Foundation for Innovation, Ontario Research Fund, and Canadian Institute for Health Research to Dr. Andrew Emili. I was supported in part by a doctoral Ontario Graduate Scholarship (2010-

2012).

First and foremost, I would like to thank and acknowledge my PhD supervisor, Andrew

Emili, for extending his support and guidance over the past 6 years. Not only did Andrew grant me the flexibility to tread into unfamiliar waters with my projects, but also served at a personal level as a great mentor and motivator. I also express my appreciation to my supervisory committee, Drs. Jeff Wrana and Dev Sidhu, for their critical insights to help keep me on track through my PhD. I also thank the many talented collaborators with whom I have had the privilege of working for their support and for taking interest in my thesis work. In particular, a special thanks to Drs. Loksum Wong and Vincent Tropepe for all things zebrafish-related; their expertise and insights were critical for applying meaningful in vivo insights to my proteomics data. I also extend my gratitude to Dr. Jack Greenblatt for his contagious enthusiasm for chromatin biology and for always being willingness to discuss all things related to transcription.

I extend my sincere gratitude all the members of the Emili Lab for ongoing collegiality and support. Last, but certainly not least, I thank my family for their patient, loving support over the past few years.

Table of Contents

Acknowledgments ...... iv

Table of Contents ...... v

List of Tables ...... ix

List of Figures ...... x

List of Appendices ...... xiii

List of Abbreviations ...... xiv

Chapter 1 Introduction ...... 1

1 Introduction ...... 2 1.1 Chromatin and its modification ...... 2 1.1.1 Histone methylation and the related protein machinery ...... 6 1.1.2 The impact of histone methylation on transcription ...... 9 1.1.3 Histone methylation, stem cells and cellular differentiation ...... 13 1.1.4 Histone methylation complexes and their importance in transcriptional regulation .... 14 1.2 The paralogous H3K9 HMTs G9a and GLP form a heterodimer required for euchromatic gene silencing ...... 20 1.2.1 Long-term gene silencing mediated by G9a/GLP ...... 23 1.3 The differentiation of retinal progenitor cells during development ...... 27 1.3.1 The origin of highly proliferative progenitor cell of the retina ...... 27 1.3.2 Regulation of RPC proliferation and cell fate decisions ...... 28 1.3.3 The role of G9a and H3K9me2 in RPCs ...... 29 1.4 Aims and objectives ...... 30

Chapter 2 A physical interaction map for histone methylation-related proteins ...... 32

2 A physical interaction map for histone methylation-related proteins ...... 33 2.1 Abstract ...... 33 2.2 Introduction ...... 34 2.3 Methods ...... 37 2.3.1 Plasmid construction ...... 37 2.3.2 Cell culture and lentivirus production ...... 37

2.3.3 Affinity purification and mass spectrometry ...... 38 2.3.4 Immunoprecipitation and western blotting ...... 39 2.3.5 Lentiviral-based shRNA-mediated mRNA knockdown ...... 40 2.3.6 Quantitative PCR ...... 40 2.4 Results ...... 40 2.4.1 A physical interaction map for histone methylation-related proteins ...... 40 2.4.2 RPRD1, RPRD1B, and RPRD2 as novel RNAPII-interacting transcriptional co-factors .. 46 2.4.3 BOD1L as a novel component of a COMPASS-like complex ...... 51 2.4.4 A novel SFMBT1-L3MBTL3 heteromeric co-repressor complex ...... 52 2.4.5 ZNF644 as a novel G9a/GLP-interacting protein ...... 53 2.4.6 ZNF644 binds G9a/GLP via an atypical C2H2-like ZF motif ...... 57 2.4.7 ZNF644 regulates global levels of H3K9me2 ...... 59 2.5 Discussion ...... 60 2.5.1 Physical interaction map of the histone methylation system ...... 60 2.5.2 The RPRD proteins bind phosphorylated CTD and possibly regulate transcription elongation and/or termination ...... 61 2.5.3 High-grade myopia protein ZNF644 physically interacts with G9a and GLP ...... 62

Chapter 3 G9a and ZNF644 physically associate to suppress progenitor gene expression and cell cycle progression in the retina ...... 64

3 G9a and ZNF644 physically associate in neuronal stem/progenitor cells to suppress progenitor gene expression and cell cycle progression in the developing retina ...... 65 3.1 Abstract ...... 65 3.2 Introduction ...... 66 3.3 Methods ...... 68 3.3.1 Zebrafish husbandry ...... 68 3.3.2 Morpholino injections ...... 68 3.3.3 Quantitative RT-PCR ...... 69 3.3.4 Whole-mount in situ hybridization ...... 70 3.3.5 Histology ...... 71 3.3.6 Retinal cross-sectional area measurement ...... 72 3.3.7 Ectopic gene expression ...... 72 3.3.8 Whole-mount antibody staining ...... 73

3.3.9 Immunohistochemistry ...... 73 3.3.10 Chromatin immunoprecipitation (ChIP) ...... 74 3.4 Results ...... 75 3.4.1 The expression of znf644 paralogues during embryogenesis ...... 75 3.4.2 Disruption of G9a or ZNF644 causes defects in the forming retina and midbrain ...... 79 3.4.3 Severe cell viability and proliferative defects are observed in g9a morphant retinas ... 84 3.4.4 Znf644a and znf644b morphant retinas both exhibit mislocalized expression of vsx2 and ccnd1 and reduced promoter H3K9me2 ...... 91 3.4.5 Znf644b morphant retinas maintain a fully multipotent proliferative state, whereas znf644a morphant retinas generate differentiated cells ...... 94 3.4.6 Summary of g9a, znf644a and znf644b morphant phenotypes in the retina ...... 101 3.4.7 H3K9me2-positive retinal nuclei are visible in differentiated cells...... 102 3.4.8 Extensive functional cooperativity between g9a and the znf644 paralogues ...... 104 3.4.9 The functions of human ZNF644 encompasses those of both zebrafish paralogues ... 107 3.4.10 The retinal defects in G9a and ZNF644 morphants are recapitulated in the midbrain 109 3.5 Discussion ...... 112 3.5.1 Neural-specific regulation of gene silencing during differentiation by G9a/ZNF644 . 112 3.5.2 Model for the role of the G9a-ZNF644 complex during RPC differentiation ...... 114 3.5.3 H3K9me2 marks differentiated neuronal cells in the retina ...... 117 3.5.4 Common cellular dependencies for G9a and ZNF644 in retinal and midbrain progenitors ...... 117 3.5.5 Possible insights into high-grade myopia ...... 119

Chapter 4 Thesis summary and future directions ...... 121

4 Thesis summary and future directions ...... 122 4.1 Thesis summary ...... 122 4.2 Future directions ...... 129 4.2.1 Characterization of novel components of histone methylation-related complexes ..... 129 4.2.2 Further tissue-specific characterization of HMT protein complex function ...... 131 4.3 Integration of HMT complex function with master transcriptional regulatory networks and upstream signalling pathways ...... 131 4.3.1 Determining the molecular factors regulating the target gene specificity of histone methylation-related complexes ...... 132

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4.3.2 How are histone methylation-related complexes related to disease processes? ...... 133

5 REFERENCES ...... 134

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List of Tables

Table 1.1: Components of human PRC2 and select functions.

Table 1.2: Components of human COMPASS complexes and their functions.

Table 1.3: Proteins known to physically associate with G9a and/or GLP.

Table 1.4: Biological roles of G9a and/or GLP during development.

Table 2.1: Primers used to clone ZNF644 isoforms into Gateway™ Entry vectors.

Table 2.2: Primers used for site-directed mutagenesis (SDM) of the predicted zinc-ion coordinating residues of ZNF644.

Table 2.3: Summary of select AP-MS data for the POLR2D and POLR2E subunits of RNAPII, as well as RPRD1A and RPRD1B.

Table 3.1: Antisense morpholino (MO) sequences used for loss-of-function investigations in zebrafish.

Table 3.2: Primer sequences used to validate MO-mediated splicing disruption of g9a, znf644a or znf644b.

Table 3.3: Primer sequences used to amplify ORFs for znf644a and znf644b for subcloning into the pCS+ vector.

Table 3.4: Primer sequences used to amplify the promoter regions of vsx2 and ccnd1.

Table 3.5: Summary of the g9a, znf644a and znf644b morphant phenotypes in the retina.

List of Figures

Figure 1.1: Schematic of the major developmental methyltransferase complexes, their enzymatic activity, and their effect on target gene transcription.

Figure 1.2: Protein domain architecture of G9a and the function(s) of these domains.

Figure 1.3: Current paradigm for sequential gene silencing by G9a/GLP-mediated H3K9me2.

Figure 2.1: Workflow outline for the lentiviral-based AP-MS approach used in this study.

Figure 2.2: Computational scoring and filtering approach for the transcription- and histone methylation-related AP-MS dataset used in this study.

Figure 2.3: Physical interaction network for histone methylation- and transcription-related protein complexes.

Figure 2.4: Hierarchal clustering of the correlation of the interaction profiles of the indicated bait proteins.

Figure 2.5: Validating interactions from AP-MS dataset using IP-WB.

Figure 2.6: The RPRD proteins specifically bind phosphorylated RNAPII.

Figure 2.7: G9a co-purified two large C2H2-like ZF-containing proteins, WIZ and ZNF644.

Figure 2.8: The atypical ZF motif of ZNF644 directly binds the G9a/GLP SET domain.

Figure 2.9: ZNF644 is a co-regulator of H3K9me2.

Figure 3.1: Protein architecture of human ZNF644 and zebrafish paralogues Znf644a and ZNF644b.

Figure 3.2: Developmental expression of znf644a and znf644b in zebrafish embryos.

Figure 3.3: Expression of znf644a and znf644b in neural progenitors of the retina and midbrain.

Figure 3.4: Disruption of g9a gives rise to severe morphological abnormalities in a dose- dependent manner.

Figure 3.5: Disruption of g9a, znf644a or znf644b yields reductions in retina size and altered midbrain morphology.

Figure 3.6: MOs targeted against znf644a or znf644b result in transcript-specific splicing defects.

Figure 3.7: Impaired cell viability in g9a morphant retinas.

Figure 3.8: G9a morphant retinas exhibit increased cellular proliferation.

Figure 3.9: G9a morphant retinas lack progenitor cells as well as various types of differentiated neurons.

Figure 3.10: Expression of vsx2 and ccnd1 in the g9a morphant retina at 48 hpf.

Figure 3.11: Znf644a and znf644b morphant retinas exhibited mislocalized expression of vsx2 and ccnd1 as well as reduced H3K9me2 levels in the vsx2 and ccnd1 promoter regions.

Figure 3.12: Persistent proliferation in znf644a and znf644b morphant retinas.

Figure 3.13: Znf644a and znf644b morphant retinas are composed of different types of cells.

Figure 3.14: Znf644a morphant retinas, but not znf644b morphant retinas, show elevated levels of apoptosis in mature populations of differentiated neurons.

Figure 3.15: Delayed differentiation in znf644b morphant embryos.

Figure 3.16: H3K9me2-positive nuclei are visible in differentiated cell populations

Figure 3.17: Extensive functional cooperativity between g9a, znf644a and znf644b.

Figure 3.18: Human ZNF644 rescues the znf644a and znf644b morphant phenotypes, but requires association with G9a/GLP.

Figure 3.19: The defects in gene expression, proliferation and survival in g9a, znf644a and znf644b morphant retinas are recapitulated in the midbrain.

Figure 3.20: Model illustrating the roles of the G9a-ZNF644 physical association in controlling multiple aspects of retinal differentiation.

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List of Appendices

Appendix 1: Spreadsheet containing the AP-MS dataset for 33 histone methylation- and transcription-related protein complexes (Worksheet 1) and the AP-MS dataset for GFP-tagged ZNF644 (Worksheet 2).

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List of Abbreviations

°C degrees Celsius aa amino acid ac acetylation ANK ankyrin AP-MS affinity purification and mass spectrometry BCIP 5-bromo-4-chloro-3'-indolyphosphate BioGRID general repository for interactions database BLAST basic local alignment tool bp base pair(s) C- carboxyl cDNA complementary DNA CID CTD-interaction domain CMZ ciliary marginal zone COMPASS complex of proteins associated with Set1 compPASS comparative proteomic analysis software suite CTD C-terminal domain DIG dioxygenin DNA deoxyribonucleic acid dNTP deoxynucleotide triphosphates ES embryonic stem Ez enhancer of zeste FITC fluorescein isothiocyanate GABA gamma aminobutyric acid GFP green fluorescent protein GST glutathione S-transferase H3K27 histone H3 lysine 27 H3K4 histone H3 lysine 4 H3K9 histone H3 lysine 9 HDAC histone deacetylase HEK human embryonic kidney HMT histone methyltransferase hpf hours post-fertilization HRP horse radish peroxidase HSPC hematopoietic stem/progenitor cells IPTG Isopropylthio-β-galactoside ISH in situ hybridization LC-MS/MS liquid chromatography and tandem mass spectrometry me1 monomethylated me2 dimethylated me2a asymmetric dimethylated me2s symmetric dimethylated xiv

me3 trimethylated mM millimolar mMO mismatch MO MO morpholino mRNA mature RNA N- amino NBT nitro blue tetrazolium chloride ncRNA noncoding RNA ng nanogram nM nanomolar NPC neural progenitor cell ORF open reading frame PAGE polyacrylamide gel electrophoresis PBS phosphate buffered saline PcG polycomb group PCR polymerase chain reaction PEV position effect variegation pg picogram PHD plant homeodomain pH3 phospho-histone H3 PKC protein kinase C PPI protein-protein interaction PRC polycomb repressor complex PTM post-translational modification RGC retinal ganglion cell RNA ribonucleic acid RNAi inhibitory ribonucleic acid RNAPII RNA polymerase II RPC retinal progenitor cell RT reverse transcription SAINT significance analysis of interactome SDS sodium dodecyl sulfate shRNA short hairpin RNA TF transcription factor Tg transgene Trx trithorax TrxG trithorax group ub ubiquitination µm micrometer µM micromolar UTR untranslated region VA versatile affinity WT wild-type ZF zinc-finger xv

Chapter 1 Introduction

1 Introduction 1.1 Chromatin and its modification

Initially defined as the “causal interactions between genes and their products, which bring the phenotype into being (Waddington, 1939), the term “epigenetics” has since evolved to describe mitotically and/or meiotically heritable changes in gene function that are not due to alterations in DNA sequence (Holliday, 1987). Epigenetics provides a powerful paradigm for the analysis of tissue and organ development in that it helps explain how the myriad cell types that comprise metazoan organisms can possess essentially the same genetic material, yet manifest cell type-specific gene expression programs that confer different proliferative and developmental potentials, unique physical properties, and characteristic attributes that persist over an extended period of time (i.e., an adult organism's lifetime).

A cell’s identity, development, behavior, and responses to its environment are defined by the set of transcription factors (TFs) that is expresses. Among the many human TFs that recognize specific DNA sequences in the context of chromatin to regulate transcription by RNAPII, certain “master” TFs are able to control cell type- specific gene expression and hence play central roles in defining cellular identity (Whyte et al., 2013). This principle is well illustrated by forcing the expression of one or more

TFs in mouse or human cells to cause transdifferentiation or the induction of pluripotency

(Zaret and Grompe, 2008; Orkin and Hochedlinger, 2011). In addition to their DNA- binding activity, TFs regulate transcription by physically interacting with various components of the transcriptional machinery as well as various chromatin-modifying and remodeling enzymes (Frietze and Farnham, 2011). The assembly and targeting of

epigenetic regulatory modules at specific chromosomal locations typically involves a complex set of physical interactions involving a host of tissue-specific TFs (Ravasi et al.,

2010), noncoding (nc)RNA molecules (Guttman et al., 2011), and auxiliary co-factor proteins. Identifying and characterizing such protein-protein interactions is critical for advancing our understanding of how gene expression patterns and cellular behavior at regulated at the molecular level.

It is now apparent that the epigenetic control of gene expression occurs broadly, if not ubiquitously, throughout embryogenesis and even throughout the life span of an organism, and that aberrant regulation thereof can cause or perpetuate disease processes.

It is also increasingly evident that epigenetic regulation of gene expression is determined in large part at the level of chromatin structure. Hence, further insights regarding the fundamental principles and processes underlying epigenetic mechanisms, in particular the composition and regulation of the associated histone-modifying machinery - and how these control chromatin structure and function to determine cellular identity, homeostatic state and dynamic behavior - are needed in order to advance our understanding of the molecular basis of tissue specification during mammalian development.

Eukaryotic DNA is organized into its characteristic “beads-on-a-string” chromatin structure by nucleosomes, which consist of 146 base-pairs (bp) of DNA intimately wrapped around an octamer of core histone proteins (two copies of each H2A, H2B, H3 and H4) and are separated by linker histones (Kornberg, 1977). Histone proteins had been known since the 1960s to undergo extensive post-translational modifications (PTMs)

(Allfrey, Faulkner and Mirsky, 1964), yet little was initially understood as to the potential impact that these marks may have on chromatin structure and function until the crystal

structure of the nucleosome was published in 1997 (Luger et al., 1997) The structural framework of the nucleosome, and the core protein components thereof, are based on two central aspects: (1) the formation of inter-histone hydrophobic interactions between internal globular domains, and (2) long, unstructured, N-terminal histone “tails” that protrude outwards from the nucleosomal core. These insights gave rise to mechanistic paradigms that are now widely accepted in chromatin biology; that covalent modification of histone tail residues, of which many comprise basic residues (i.e., lysine and arginine), can alter chromatin packing and, hence, the accessibility of DNA, either by altering internucleosomal interactions or by facilitating the recruitment of specific chromatin- modifying (enzymatic) complexes via the molecular recognition of and site-specific binding to modified histone tails (Bannister and Kouzarides, 2011). It rapidly emerged that histone-modifying enzymes, including histone acetyltransferases (HATs) and protein kinases, can alter the intrinsic stability of nucleosomes and influence the compaction of chromatin. In addition, various histone PTMs, either alone or in combination, have been posited to form a “histone code” (Kouzarides, 2007; Taverna et al., 2007) that is “read” by various effector proteins and protein complexes that associate specifically with the modified nucleosomes (Taverna et al., 2007). These histone PTMs and their dynamic regulation in the wake of physiological cues and developmental signals are now seen as a common unifying theme that underlies essentially all DNA-templated processes, including gene transcription, chromosome compaction, and DNA replication and repair.

The many covalent histone PTMs that are known to occur on histone proteins include the following: Lysine acetylation, lysine and arginine methylation, serine and threonine phosphorylation, ubiquitination, sumolyation, ADP-ribosylation, arginine

deimination (to citrulline) and proline isomerization. These PTMs are known to affect at least two-thirds of the potentially modifiable amino acid residues in the four core histones

(Kouzarides, 2007). Many of the enzymes that carry out site-specific histone PTMs have been identified in certain cases, although it should be noted that histone-modifying enzymes might also target non-histone substrates for methylation, thus enabling a distinct, and perhaps complementary, mode of action. Particular chromatin marks have also been shown by chromatin immunoprecipitation (ChIP)-chip and ChIP-sequencing studies to exhibit loci specific genome-wide patterns that often correlate with gene expression and specific biological phenomena (Pokholok et al., 2005; Liu et al., 2005).

Moreover, these marks are generally reversible (e.g., acetylation by histone deacetylases

[HDACs], phosphorylation by phosphatases, methylation by demethylases, and ubiquitination by deubiquitinases), thus providing a counter-balance that enables more flexibility in controlling gene expression during development. The enzymes that create or remove these chromatin marks are generally thought to be regulated by the presence or absence of other marks on the same or adjacent histones (Fischle et al., 2005; Nelson,

Santos-Rosa and Kouzarides, 2006; Kouzarides, 2007), and, in some (and perhaps most) cases, it is the local configuration of histone PTMs, which can be recognized by multiple domains within one or more associated effector proteins or complexes, that is interpreted by the downstream epigenetic "reader" machinery to regulate a particular process (Fischle et al., 2005). These effector chromatin-related protein complexes then either perform additional essential chromatin alterations or else directly control various chromatin-based processes (e.g., transcription or heterochromatin formation).

1.1.1 Histone methylation and the related protein machinery

Notable among the constellation of conserved histone tail modifications

(Kouzarides, 2007), is methylation of ε-amino groups of lysine and ω-guanidine groups of arginine by histone methyltransferase (HMT) enzymes. Relative to other histone

PTMs, lysine and arginine methylation comprise a higher degree of structural complexity in that these basic side chains can accept multiple methyl groups. Specifically, lysine residues can be monomethylated (me1), dimethylated (me2) or trimethylated (me3), while arginine residues can be monomethylated, symmetrically dimethylated (me2s) or asymmetrically dimethylated (me2a). In contrast to PTMs that neutralize or induce changes in intrinsic histone charge state (e.g., lysine acetylation by lysine acetyltransferases and serine phosphorylation by protein kinases, respectively), lysine or arginine methylation increases the hydrophobic character of the modified side chain without affecting overall charge. In terms of biological significance, site-specific lysine methylation marks have been linked to the establishment and/or maintenance of specific chromatin states, such as heterochromatin formation (H3K9me3, H3K27me3,

H4K20me3), broad transcriptionally active (e.g., H3K4me3, H3K36me3) and silent

(H3K9me2/me3 and H3K27me3) genomic regions, and localized sites of DNA repair

(H3K79me2, H4K20me2) (Martin and Zhang, 2005), to name a few. The corresponding

HMTs – broadly viewed as the so-called “writers” of these histone methylation marks – often exhibit specificity both in terms of the degree of methylation they generate as well as their cognate substrate recognition sequences (e.g., surrounding amino acid sequence of targeted residues they modify).

The discovery of HMTs has roots in classic genetic screens in Drosophila aimed at identifying regulators of the position effect variegation (PEV) phenomena, wherein genes inserted near heterochromatin were found to be silenced, leading to the discovery of both suppressors and enhancers of variegation (Su(var) and E(var) genes, respectively)

(Reuter and Spierer, 1992). Subsequent genetic studies in Drosophila identified two additional families of proteins, the Trithorax group (TrxG) and the Polycomb group

(PcG) of regulators, as playing a central role in the control of gene expression throughout development (Ringrose and Paro, 2004). Many of the genes identified in such screens

(e.g. Suppressor of variegation 3-9 (Su(var)3-9, Enhancer of zeste (Ez) and Trithorax

(Trx)) encoded proteins with a common 130 amino acid motif known as the SET domain

(Jenuwein et al., 1998). In 2000, it was demonstrated that the SET domain of Drosophila

Suv39h (and its homologs) possesses intrinsic site-specific HMT activity toward H3K9

(Rea et al., 2000). Since then, extensive structure-function analyses have been applied to a number of proteins harboring SET domains or possessing lysine and arginine HMT activity, collectively encompassing a set of some 60 putative enzymes encoded by the human genome. Most of these enzymes appear to function as components of multi- subunit complexes, and a sizeable number of macromolecular assemblies containing

HMTs as integral components have been identified in diverse eukaryotic organisms, some of which have been implicated in directing gene expression and cell fate decisions in different physiological contexts.

Not only is the activity of HMTs generally reversible by histone demethylase enzymes – the “erasers” of histone methylation – but also the site-specific catalytic activity of HMTs is often complemented by highly evolved and expanded families of

methyl-lysine and methyl-arginine recognition domains. These so-called “readers” of histone methylation are often found as components of larger multiprotein effector complexes. Following the discovery that the chromodomain (chromatin organization modifier) of heterochromatin protein 1 (HP1) binds H3K9me3 (Bannister et al., 2001), several sequence- and structurally-related modules capable of recognizing other histone methylation marks were discovered. These include specific recognition by plant homeodomain (PHD) fingers and WD40 repeats, as well as by various modules of the

Royal superfamily (which comprises Tudor domains, chromodomains, MBT, and PWWP domains) (Maurer-Stroh et al., 2003). These domains are highly expanded from single cell yeast to metazoa: the PHD domain is 5-fold more frequent in humans relative to S. cerevisiae, the Royal family (Chromo, TUDOR, PWWP and MBT) 9-14-fold, and the

MBT domain (prominent in metazoans) is absent in yeast (On et al., 2010). This striking enhancement in the number and variety of domains that write and read histone methylation in metazoans likely reflects the greater need for fine-tuned regulation of transcription and other chromatin-bases processes that are essential for cell-type-specific gene expression and tissue-specific development.

Histone methylation is now among the better-characterized epigenetic marks known to enable the establishment and maintenance of precise cell- and tissue-specific gene expression programs that are essential for proper metazoan development. In addition, the corresponding enzymes represent an emerging class of clinically relevant drug targets (particularly in cancer) (Copeland, Moyer and Richon, 2013; Copeland,

Solomon and Richon, 2009; Arrowsmith et al., 2012). Moreover, over the past decade, important discoveries and rapid technological advances have impacted our understanding

of the dynamic landscape of mammalian epigenomes, how the epigenome is shaped by histone methylation-related complexes, and how deregulation of these complexes and systems can induce developmental defects or underlie disease processes. As the functional significance of key multicomponent HMT complexes in the regulation of gene expression programs continue to come to light, more detailed patterns are emerging regarding the epigenetic underpinnings of differential gene expression programs and their impact on cell fate decisions. The opportunity to use an unbiased proteomic strategy to explore the molecular machinery governing histone methylation (and associated chromatin modifications) and to subsequently investigate how these physical and regulatory interactions instruct developmental processes motivated my thesis studies in the Emili laboratory.

1.1.2 The impact of histone methylation on transcription

The initial connection between histone methylation and active transcription within euchromatin was discovered in the ciliated protozoan Tetrahymena, where H3K4 methylation was observed in the transcriptionally active macronucleus, but not in the silent micronucleus (Strahl et al., 1999). The importance of H3K4 methylation, as well as many other alternate histone methylation marks, to transcription is reflected in recent

ChIP-chip and ongoing ChIP-sequencing studies that have revealed genome-wide patterns of methylation marks correlating with transcriptional states and the presence of cis-regulatory elements, including enhancer and promoter regions, exons and introns, and various phases of the transcription cycle (e.g. initiation, elongation) (Black, Van and

Whetstine, 2012). Certain aspects regarding the influence of histone methylation in the regulation of transcription, particularly RNA stalling and the processivity of RNA

polymerase II (RNAPII), have been well studied (described below), although many mechanistic details remain unresolved. A summary of key HMT complexes, their core compositions and enzymatic activities, and their effects on the transcriptional output of their target genes is provided in Figure 1.1. The identification of factors that regulate and mediate the recruitment of these and other HMT complexes to their respective genomic regions is currently an area of intense investigation.

Figure 1.1: Schematic representation of the major developmental HMT complexes, their substrate specificity, and their effect on target gene transcription. (Left) The core composition of COMPASS, PRC2 and G9a/GLP protein complexes, with enzymatic and lysine residue of histone H3 highlighted. (Right) Positioning of histone methylation mark within gene bodies and the corresponding effect on target gene transcription.

1.1.2.1 Active transcription

Active genes are characterized by histone hyperacetylation (e.g., on various lysine residues of the N-terminal tails of histones H3 and H4) in their promoter regions, as well as trimethylation on H3K4, H3K36 and H3K79, and ubiquitination on H2BK120. The mechanisms leading to the generation of H3K4me3, H3K36me3 and H3K79me3 are co- transcriptional – in other words, the HMTs responsible for these modifications physically associate with RNAPII, usually via the C-terminal domain (CTD) of its largest subunit,

RPB1. Unique among eukaryotic RNA polymerases, the CTD of RNAPII contains multiple, sometimes degenerate, repeats (52 in humans) of the heptapeptide sequence

YSPTSPS. The CTD is important for transcription initiation, elongation and termination, and for the coupling of transcription to histone modification and various aspects of mRNA processing (e.g. capping, splicing, polyadenylation) (Buratowski, 2009). The

CTD repeats can be phosphorylated at S2, S5 and S7 (Kim et al., 2009; Komarnitsky,

Cho and Buratowski, 2000), and each these phosphorylation events is closely coupled with histone methylation in the transcribed region. For example, phosphorylation of repeat residue S5 (S5P) is critical for targeting the COMPASS HMT complex, which generates H3K4me2/me3 at promoter regions (Shilatifard, 2012). Once RNAPII escapes from the promoter region during transcription initiation, another HMT DOT1L, which resides in a “super elongation complex” that also contains the CTD S2 kinase P-TEFb

(Luo, Lin and Shilatifard, 2012), generates H3K79me3 marks. In addition, the HMT

SET2 recognizes phosphorylation of the CTD at S2 (S2P), leading to the downstream formation of H3K36me3 during transcription elongation (Krogan et al., 2003). In yeast, this H3K36 methylation mark is recognized by the Rpd3S histone deacetylation complex,

which assembles deacetylated nucleosomes in the wake of elongating RNAPII, preventing cryptic transcription initiation (Carrozza et al., 2005). Hence, physical interactions between RNAPII and HMTs occur frequently within a gene body, resulting in the establishment of differential histone methylation marks.

1.1.2.2 Repressed transcription

In contrast to the histone PTMs that are observed at actively transcribed genes, transcriptionally “silent” (i.e., transcriptionally inactive or repressed) chromatin is characterized by high levels of methylation on either H3K9 or H3K27, concomitant with

DNA methylation and generally low levels of histone acetylation. The repressive H3K27 methylation mark is generated by the multi-component Polycomb repressive complex

(PRC)2 (Schwartz and Pirrotta, 2008), which has been extensively studied. H3K9 methylation is generated by several different HMTs, but gene silencing in euchromatic regions is largely controlled by the paralogous HMTs G9a and GLP, which form a stoichiometric heterodimer in vivo (Shinkai and Tachibana, 2011). Recent findings suggest cross-talk among these repressor complexes. For example, H3K9me2 by

G9a/GLP can regulate gene silencing by PRC2 in certain cellular contexts (Mozzetta et al., 2013). Unlike gene activation, which is generally thought to be targeted to specific loci by site-specific DNA-binding proteins (i.e., TFs), the process of gene silencing is sometimes (if not usually) targeted to particular loci by protein complexes containing non-coding RNAs (ncRNAs) as integral components (Lee, 2012). For example, small ncRNAs, such as miRNAs, piRNAs and siRNAs, generated from larger precursor transcripts, are incorporated into RITS (RNA-induced transcription silencing) complexes

(Verdel et al., 2004), which are thought to be recruited to nascent transcripts via base-

pairing interactions (Bühler, Verdel and Moazed, 2006), leading to polycomb-dependent silencing (Kim et al., 2006). Nonetheless, much work is required to fully understand the suite of factors and the seemingly elaborate molecular mechanisms that orchestrate the recruitment of repressive HMT complexes to their target gene loci.

1.1.3 Histone methylation, stem cells and cellular differentiation

In embryonic stem (ES) cells, pluripotency-related TFs establish a regulatory network that creates and maintains a pluripotent state characterized by extensive open chromatin and relatively little heterochromatin (Meshorer and Misteli, 2006). This principle is well illustrated during somatic cell reprogramming, where the forced expression of a set of pluripotency-related TFs can revert somatic cells back to a pluripotent-like epigenetic state (Takahashi and Yamanaka, 2006). These TFs and other auxiliary factors are thought to recruit multiple chromatin-modifying complexes (Dejosez et al., 2008) to establish a local cooperative feedback network of both positive and negative regulation, ultimately stabilizing the pluripotent state (Boyer et al., 2005). There is now mounting evidence that establishing and maintaining the pluripotent state, as well as exiting pluripotency to differentiated states, involve extensive histone methylation. For example, maintaining a pluripotent state requires the repressed expression of a large cohort of developmental genes, which seems to be achieved, at least in part, by interactions between the pluripotency TFs and the HMT complexes PRC2 and PRC1, which generate and recognize, respectively, the H3K27me3 silencing mark

(Schuettengruber et al., 2007). Moreover, efficiently creating pluripotent cells and even bypassing the need for certain pluripotency-related TFs can be accomplished by treating certain somatic cells with an inhibitor specific for the H3K9 HMTs G9a/GLP (Kubicek et

al., 2007), suggesting that a transient reversal of gene silencing is also important for induced pluripotency. In addition, an alternative route to reprogramming is via the fusion of somatic cells into an ES cell environment, which can be enhanced by reducing G9a activity (Ma et al., 2008). Certain ncRNAs seem play important roles in controlling the expression of pluripotency genes through physical associations with HMT complexes.

For example, the expression patterns of multiple ncRNAs correlate with ES cell pluripotency or differentiation (Dinger et al., 2008), and some of these can bind to chromatin-modifying complexes that repress (e.g. PRC2) or activate (e.g. COMPASS) transcription (Guttman et al., 2011). However, considerable work still needs to be done to reveal the molecular mechanisms driving the assembly, targeting and function of these and other HMT complexes - both in ES cells, and also in various other types of tissue- specific progenitor cell populations.

1.1.4 Histone methylation complexes and their importance in transcriptional regulation

Like other proteins that bear enzymatic, structural or regulatory properties, histone methylation-related enzymes typically operate as parts of modular ‘molecular machines’ linked together by specific physical interactions. Understanding the core subunit composition, abundance, regulation and functional relationships of these complexes is therefore essential to understand how histone methylation is regulated during development. Indeed, the importance of protein complex formation for conferring target gene specificity and determining the enzymatic activity of HMT complexes has been amply illustrated in PcG and TrxG protein complexes, which I will briefly describe below.

1.1.4.1 Polycomb repressor complexes, H3K27 methylation, and transcriptional repression

PcG and TrxG proteins operate biochemically as distinct, yet evolutionary conserved enzymatic complexes that, together, ensure proper spatial and temporal expression of master regulator TFs genes during development (Schuettengruber et al.,

2007). For example, Hox genes are situated contiguously as clusters and encode homeodomain TFs that regulate diverse cellular signaling pathways involved in development and disease (Shah and Sukumar, 2010). In addition to Hox genes, TrxG proteins also regulate the expression of many other genes that are important for patterning, cell proliferation, and stem cell identity by maintaining genes in an active state (Fisher and Fisher, 2011). The HMT complex PRC2 is the only complex known to regulate the repressive H3K27 methylation mark. Several aspects of PRC2 function remain under intense investigation, including the exact composition and functions of mammalian PcG complexes (in light of a vastly expanded number of paralogue subunits), the physical associations that direct and specify PcG to particular loci to facilitate chromosomal reorganization, and the mechanisms whereby PcG complexes prevent elongation by RNAPII at target genes.

PRC2 is composed of four subunits: EZH2/EZH1, SUZ12, EED and

RBBP4/RBBP7. The inner workings of these core components and their contribution to

PRC2 function have been extensively characterized (O'Meara and Simon, 2012). EZH1 and EZH2 are paralogous HMTs that harbor catalytic SET domains, yet each forms mutually exclusive PRC2 complexes that depend critically on physical inputs from other core PRC2 subunits (particularly EED and SUZ12) for maximal catalytic activity

(O'Meara and Simon, 2012). The EED subunit functions as an allosteric regulator of

PRC2 and can stimulate enzymatic activity through direct recognition of H3K27me3 peptides (Xu et al., 2010), perhaps providing a means for propagating the H3K27me3 onto neighboring nucleosomes or during DNA replication (Ahn, Keogh and Buratowski,

2009). The subunit SUZ12 mediates PRC2 complex assembly and likewise acts as an allosteric activator of EZH2 enzymatic activity (Cao and Zhang, 2004). Additional complex components also regulate PRC2 activity, including the zinc-finger protein

AEBP2, the multiple PCL homologs (PCL1, PCL2 and PCL3) and JARID2 (Margueron and Reinberg, 2011). Table 1.1 below contains a brief summary of PRC2 components and their putative functions.

The expansion of PRC2 complex components (and various PcG components in general) in mammalian systems is thought to enable the assembly of more specialized, compositionally diverse complexes of related functionality. In the case of PRC2, EZH1 and EZH2 (which form mutually exclusive PRC2 complexes of similar global composition) show functional redundancy in ES cells (Shen et al., 2008) and hair follicle homeostasis and wound repair (Ezhkova et al., 2011). Yet EZH1 and EZH2 also function non-redundantly in other cellular contexts, such as during myogenic differentiation where

EZH2-PRC2 strongly co-localizes with H3K27me3 and EZH1-PCR2 with H3K4me3 marked chromatin (Mousavi et al., 2012). Interestingly, EZH1-PRC2 plays a role in stimulating transcription elongation in this context, though the mechanistic basis of this regulation and why EZH1, but not EZH2, is used preferentially remains unclear.

Understanding the diversity in composition, regulation and significance of the different mammalian PRC2 complexes is an important and active area of research interest.

1.1.4.2 COMPASS-related protein complexes, H3K4 methylation, and transcriptional activation

Most TrxG proteins exert their function as part of large multi-protein complexes that regulate transcription via histone methylation or chromatin-remodeling. The current understanding of TrxG HMT complexes mainly stems from biochemical purification and functional characterization of the yeast H3K4 methyltransferase Set1. In budding yeast,

Set1 resides in an 8-member complex known as COMPASS (Complex Proteins

Associated with Set1), of which the other core COMPASS subunits (e.g. Cps50 and

Cps30) are known to be essential for catalytic activity (Shilatifard, 2012; Cosgrove and

Patel, 2010). COMPASS is recruited to initiated or promoter-proximal paused forms of

RNAPII through direct recognition of the phosphorylated RNAPII CTD at S5 (CTD-S5) by the Cps35/Swd2 subunit in yeast, as well as the orthologous WDR82 subunit of related mammalian COMPASS-like complex (Lee and Skalnik, 2008).

Table 1.1. Components of human PRC2 and select functions.

Subunit Function EZH1/EZH2 Generation of H3K27me2/me3 Allosteric activator of methyltransferase activity; Binding SUZ12 H3K4me3 or H3K36me3 peptides reduces activation Binds H3K27me3 peptides; Allosteric activator of EED methyltransferase activity RBBP4/RBBP7 Histone chaperones; Binds unmodified residues 1-10 of H3 PCL1/PCL2/PCL3 Binds H3K36me3 peptides via tudor domains Allosteric regulator of methyltransferase activity; Targeting to AEBP2 specific DNA sites Co-recruited with PRC2 to target genes; Possible targeting JARID2 factor (CG rich)

In contrast to a single type of COMPASS complex expressed in yeast cells, human cells express at least six orthologues of Set1 (SETD1A, SETD1B, MLL1, MLL2,

MLL3 and MLL4) that each form a distinct COMPASS-like complex. Each of these related COMPASS-like complexes contains at its core a Set1 orthologue as well as four additional core cofactors (ASH2L, RBBP5, WDR5 and DPY30) that have yeast orthologues. The human Set1 orthologues are much larger in size and contain a more diverse array of protein domains that impart unique functional properties. In addition to the common four core subunits, each COMPASS-like complex contains unique subunits.

For instance, SETD1A and SETD1B, which are most closely related to yeast Set1, contain the yeast CTD S5P-binding subunit WDR82, which mediates recruitment (and hence H3K4 HMT enzymatic activity) to S5 phosphorylated CTD (Lee and Skalnik,

2008). SETD1A and SETD1B versions of COMPASS-like complexes also uniquely contain the subunit CXXC1, which binds to unmethylated CG-rich DNA regions (known as CpG islands), thereby regulating the global positioning of H3K4me3 in ES cells

(Clouaire et al., 2012). Conversely, the MLL1- or MLL4-containing COMPASS-like complexes, which are most closely related to Drosophila Trx protein, uniquely contain the tumor suppressor MENIN, which is implicated in targeting MLL1 to the promoters of

Hox gene loci during hematopoiesis (Hughes et al., 2004). MLL3- and MLL4-containing

COMPASS-like complexes contain the H3K27 demethylase KDM6A/UTX, which likely helps to reverse gene silencing by PRC2 at gene loci (Agger et al., 2007). A summary of annotated COMPASS-like complex components and their reported functions is found in

Table 1.2 below.

Table 1.2. Common and subtype-specific components of human SET1/MLL complexes and select functions.

Subunit SET1/MLL complex Function SETD1A/SETD1B, ASH2L MLL1/MLL2, Stimulates methyltransferase activity MLL3/MLL4 SETD1A/SETD1B, RBBP5 MLL1/MLL2, Stimulates methyltransferase activity MLL3/MLL4 SETD1A/SETD1B, Stimulates methyltransferase activity; WDR5 MLL1/MLL2, Binds unmethylated and methylated MLL3/MLL4 H3K4 peptides SETD1A/SETD1B, DPY30 MLL1/MLL2, Stimulates methyltransferase activity MLL3/MLL4 WDR82 SETD1A/SETD1B Binds S5 phosphorylated RNAPII CXXC1 SETD1A/SETD1B Binds unmethylated CpG islands MENIN MLL1/MLL4 Targets MLL1 to Hox loci HCF1 MLL2/MLL3 UTX MLL2/MLL3 H3K27 demethylase PTIP MLL2/MLL3 PA1 MLL2/MLL3 NCOA6 MLL2/MLL3 Nuclear receptor coactivator

Understanding the unique composition of human COMPASS-like complexes has provided critical insights regarding the differential functions of these complexes in transcription, development and disease. This is particularly true for MLL1 because reciprocal chromosomal translocations involving MLL1 and a variety of fusion partners have been causally linked to human myeloid malignancies such as acute myeloid and lymphoid leukemias (Marschalek, 2011). The C-terminal region harbors the SET domain as well as a binding interface for the other core COMPASS subunits, whereas the N- terminal fragment is responsible for targeting MLL1 activity to Hox loci via physical

association with targeting co-factors, such as MENIN (Hughes et al., 2004). As with

PcG-related protein complexes, understanding how COMPASS-related complexes are regulated and how compositional differences among complexes modulate complex function remain largely unknown.

1.2 The paralogous H3K9 HMTs G9a and GLP form a heterodimer required for euchromatic gene silencing

Following the discovery of SUV39H1 as the first H3K9 HMT, G9a was reported as a second H3K9 HMT that also exhibited more limited activity toward H1 and H3K27 in vitro (Tachibana et al., 2001). It was more recently reported that G9a can also methylate histone H3 at lysine 56, which seems to be important for loading of the DNA replication factor PCNA during S-phase (Collins et al., 2005). Although both SUV39H1 and G9a preferentially catalyze mono-, di- and tri-methylation of H3K9 in vitro (Collins et al., 2005; Kubicek et al., 2007), their cellular activities are non-overlapping in vivo because mouse ES cells deficient in Suv39h1 and Suv39h2 only lack the H3K9me3 mark at pericentric heterochromatin, while G9a deficient cells lose H3K9me1 and H3K9me2 marks in euchromatic regions (Tachibana et al., 2001; Peters et al., 2003; Rice et al.,

2003). The additional G9a paralogue, known as GLP (G9a-like protein), harbors a nearly identical SET domain in its C-terminus (Figure 1.2), possesses equivalent HMT activity and substrate specificity in vitro (Ogawa et al., 2002; Tachibana et al., 2005; Weiss et al.,

2010) and, like G9a, is required for H3K9me1 and H3K9me2 in euchromatic regions

(Tachibana et al., 2005). In virtually all cell types investigated, G9a and GLP form a stoichiometric heteromeric complex via their SET domains (Ueda et al., 2006). Thus,

G9a and GLP are functionally overlapping and non-redundant.

Chromatin H3K9me2 marks strongly correlate with transcriptionally inactive regions of the genome (Barski et al., 2007), occupying both isolated genomic regions as well as larger megabase-long blocks of chromatin (Wen et al., 2009; Chen et al., 2012), where they are thought to serve structural roles in maintaining epigenetic memory during lineage formation. The ability of G9a/GLP to generate large blocks of H3K9me2 marked chromatin may be due to their ability to both write and read the H3K9me1 and H3K9me2 marks (Collins and Cheng, 2010), the latter capability owing to the presence of ankryin

(ANK) repeats which can bind to both H3K9me1 and H3K9me2 (Collins et al., 2008).

However, how the H3K9me2 mark is initially generated and maintained in vivo, as well as the tissue-specific cofactors that likely determine the lineage-specific patterning of

H3K9me2, are currently unknown.

Figure 1.2: Protein domain organization of the H3K9 methyltransferase G9a (and

GLP). (Top) Crystal structures of the ankryin repeats, which bind H3K9me1 and

H3K9me2 peptides, and the enzymatic SET domain, which generates the H3K9me1 and

H3K9e2 marks. Adapted from (Collins and Cheng, 2010).

G9a/GLP preferentially methylate “naked” histone octamers (as opposed to nucleosomal histones) in vitro (Tachibana et al., 2005; Shinkai, 2007). Together with their co-localization at DNA replication foci during S-phase (Esteve et al., 2006), it has been proposed that G9a/GLP target newly synthesized histones for methylation immediately prior to their insertion into chromatin. Nonetheless, given the absence of an obvious DNA binding domain, the recruitment of G9a/GLP to appropriate target loci

likely requires physical association with one or more sequence-specific TFs and/or ncRNAs. Several G9a/GLP-interacting proteins, as well as some putatively associated ncRNA molecules, have been reported (Table 1.3). In ES cells, the G9a/GLP heteromeric complex physically associates with a large C2H2 ZF motif-containing protein known as

WIZ (Ueda et al., 2006). Endogenous levels of GLP and WIZ are required maintain the

G9a protein stability, and WIZ is important for bridging the HMTs to the CtBP co- repressor machinery (Ueda et al., 2006), which functions in concert with DNA-binding repressors (such as ZEB) with histone deacetylation and histone demethylation enzymes

(Shi et al., 2003). However, how these and other G9a/GLP interacting proteins regulate the activity of G9a/GLP in various tissues during development is generally unknown.

1.2.1 Long-term gene silencing mediated by G9a/GLP

G9a and H3K9me2 have been associated with the transcriptional repression of diverse genes in different cellular contexts: for example, silencing of the Oct4 gene in differentiating ES cells (Feldman et al., 2006), in NSRF/REST-mediated silencing of neuronal genes in non-neuronal lineages (Roopra et al., 2004), and in PRDI-BF1- mediated silencing during B-cell differentiation (Gyory et al., 2004). The regulatory roles of G9a/GLP in diverse physiological contexts are summarized in Table 1.4.

Table 1.3. Proteins known to physically interact with G9a and/or GLP.

Protein Function Reference CDP/Cut Transcriptional repression (Nishio and Walsh, 2004) E2F6 Transcriptional repression (Ogawa, 2002) Gfi1 Transcriptional repression (Duan et al., 2005) (Vassen, Fiolka and Möröy, Gfib Transcriptional regulation 2006) PRDM1 Transcriptional repression (Gyory et al., 2004) NRSF/REST Transcriptional repression (Roopra et al., 2004) PRDM6 Transcriptional repression (Davis et al., 2006) ZNF217 Transcriptional repression (Banck et al., 2009) UHRF DNA methylation (Kim et al., 2009) Heterochromatin; HP1 transcriptional repression; (Nozawa et al., 2010) methyl-lysine binding DNMT1 DNA methylation (Esteve et al., 2006) (Epsztejn-Litman et al., DNMT3A/DNMT3B DNA methylation 2008) WIZ Unknown (Ueda et al., 2006) ZNF200 Unknown (Nishida et al., 2007)

Table 1.4. Biological roles for G9a and/or GLP during development.

Organism Context Function Reference(s) Drosophila Embryogenesis Learning and memory deficits (Kramer et al., 2011)

Zebrafish Embryogenesis Defects in neurogenesis (Rai et al., 2010) Silencing of pluripotency- (Nagano et al., Mouse Embryogenesis associated loci; Silence imprinted 2008; Wagschal et al.,

genes 2008) Defective chromosome Mouse Germ line (Tachibana et al., 2007) segregation Mouse Lymphocytes Impaired immune response (Thomas et al., 2008) Mouse B-cell Impaired immune response (Thomas et al., 2008) Mouse T-cell Impaired cytokine expression (Lehnertz et al., 2010) De-repression of neuronal Mouse Postnatal forebrain progenitor genes; defects in (Schaefer et al., 2009) cognition and adaptive behaviors Silencing of non-cardiac genes;

Mouse Cardiomyocytes regulation of morphogenesis of (Inagawa et al., 2013) atrioventricular septum Hematopoietic Control the timing of lineage Mouse (Chen et al., 2012) stem/progenitor cells commitment and differentiation Reduced locomotor activity and (Schaefer et al., Mouse Brain exploration; increased anxiety and 2009; Balemans et al., altered social behavior 2010)

The silencing of the Oct4 gene – as well as other genes necessary for preserving

pluripotency – serves as a commonly held model for the step-wise manner in which target

genes are silenced by G9a/GLP (Epsztejn-Litman et al., 2008; Feldman et al., 2006)

(Figure 1.3): (1) transcriptional inactivation by transiently expressed repressor factors;

(2) HDAC-mediated removal of activating histone acetylation marks and subsequent

G9a/GLP-mediated methylation of H3K9; (3) recruitment of the heterochromatin protein

HP1, leading to the formation of local heterochromatin; and (4) subsequent de novo DNA

methylation by DNMT3A and DNMT3B. Both G9a and GLP are essential for the long-

term silencing of Oct4 and other pluripotency-associated genes, and ES cells lacking G9a or GLP can be readily reverted to a pluripotent state, even after differentiation has been initiated (Feldman et al., 2006). Thus, G9a/GLP-mediated silencing, in collaboration with

HDACs and DNMTs, represents a highly stable long-term form of transcriptional inactivation.

Figure 1.3: Current model for gene silencing by G9a/GLP-mediated H3K9me2.

Changes in histone modifications required for the suppression of pluripotency/multipotency- and proliferation-related genes during cellular differentiation. In the case of the Oct4 gene locus during ES cell differentiation, a G9a- containing enzymatic complex is required for the removal of activating histone methylation and acetylation marks and the concomitant deposition of repressive histone and DNA methylation marks. Adapted from (Cedar and Bergman, 2009).

1.3 The differentiation of retinal progenitor cells during development

Epigenetic mechanisms have proven to be critical regulators of development in diverse physiological contexts, as illustrated during the formation of the central nervous system (CNS). Aberrant brain function can be caused by defects in chromatin PTMs, and histone methylation-related proteins have been linked to normal physiological neural plasticity and multiple forms of behavioral memory (Dulac, 2010; Borrelli et al.,

2008; Levenson and Sweatt, 2005). Recent studies have shed light on the importance of proper regulation of chromatin regulatory complexes in CNS development and disease, although focus has been placed mostly on TrxG and PcG complex proteins during adult neurogenesis (Lessard and Crabtree, 2010; Yang et al., 2012; Ma et al., 2010). For the purposes of my thesis, I will briefly describe an overview of the current state of knowledge with respect to the epigenetic regulation of cell proliferation and differentiation programs focusing on the developing retina as a relevant system.

1.3.1 The origin of highly proliferative progenitor cell of the retina

Retinal cells are initially derived during embryogenesis from neuroectoderm cells from within the prosencephalon region of the developing neural tube (Gilbert SF, Gilbert and Knisely, 2006). After gastrulation (where the three germ layers are formed) has taken place, inductive extracellular signals emanating from the notochord trigger the formation of the neural tube from a population of overlying ectodermal cells. It this this neural tube structure that (via non-uniform cellular proliferation) engenders the three primary brain vesicles: the forebrain, midbrain, and hindbrain. Extracellular signalling pathways are responsible for inducing the formation of the presumptive eye field in the forebrain

region (Sernagor, 2006), but it is the expression of TFs, such as Rx1, that induces neuroepithelial cells of the eye field to undergo bilateral evagination, thus producing the optic vesicle (Mathers et al., 1997). Rx1 activates the expression of other eye field TFs, such as Pax6 and Six2, that stimulate retinoblast proliferation (Zuber et al., 2003; Zhang,

Mathers and Jamrich, 2000). The cooperative activity of these eye field TFs results in the invagination of the optic vesicle to produce the bilayered optic cup composed of the neuroblastic layer, which will then give rise to RPCs, and the retinal pigment epithelium.

1.3.2 Regulation of RPC proliferation and cell fate decisions

Early RPCs in the newly formed optic cup initially and uniformly divide symmetrically – that is, give rise to two proliferative daughter cells during cell division – thereby increasing the progenitor population and expanding the tissue. The maintenance of a proliferative multipotent state requires the upregulated expression of multipotency- related TFs, such as Pax6, Vsx2 and Sox2. Accordingly, it is well-documented that mutations in these genes are linked to proliferative defects characterized by small or absent eyes (Levine and Green, 2004; Green, Stubbs and Levine, 2003; Taranova et al.,

2006). Later in retinal development, RPCs begin to display considerable heterogeneity, both in their patterns of cell division and gene expression (Trimarchi, Stadler and Cepko,

2008; Dyer and Cepko, 2001). Despite this apparent heterogeneity among RPCs, these cells as a population undergo unidirectional changes in their competence and begin to withdrawal from the cell cycle and give rise to differentiated progeny of specialized function (in a birth order that appears to be evolutionary conserved), thus producing the seven major classes of retinal cell types found in the adult retina (La, Rapaport and

Rakic, 1991; Rapaport et al., 2004). Importantly, the proper differentiation of retinal cells

(and hence a properly formed and functional retina) critically depends upon the tight coordination of RPC proliferation and differentiation – as well as with eye growth – to ensure that the forming retina achieves its proper size, morphology and structure (Green,

Stubbs and Levine, 2003; Wong et al., 2010). Indeed, defects in this coordination are know to cause a small eye phenotype known as microphthalmia (Levine and Green,

2004) as well as a propensity to retinal degeneration and blindness. In proliferating RPCs, master multipotency-related TFs (such as Pax6, Vsx2 and Sox2) are thought to maintain a proliferative multipotent state by establishing an RPC-specific gene regulatory network.

Chief among these, Vsx2 maintains a proliferative multipotent state by repressing the expression of negative regulators of the cell cycle (i.e., p27kip1) (Wong, Conger and

Burmeister, 2006) as well as pro-neural TFs factors that restrict lineage potential (i.e.,

Ath5, Foxd4, Vsx1, etc.) (Vitorino et al., 2009). However, very little is known regarding the cell intrinsic molecular machinery responsible for the proper integration of cellular proliferation, reduced cellular competency, cell cycle withdrawal and differentiation, thus ensuring the efficient and accurate generation of each cell type at the appropriate stage during development.

1.3.3 The role of G9a and H3K9me2 in RPCs

At the time I began working on this project, virtually nothing was known about the potential role of G9a/GLP in the retina. It has since been demonstrated in zebrafish that G9a functions cooperative with DNMTs to regulate the development of several physiological systems, including formation of the CNS (Rai et al., 2010). In 2012, Katoh et al. (Katoh et al., 2012) characterized an essential role of G9a in mouse RPCs. Among the defects they observed upon RPC-specific deletion of G9a were elevated levels of

apoptosis, partial loss of the outer nuclear layer, and persistent cellular proliferation

(Katoh et al., 2012). RPC-specific genes, including multipotency-related (TFs Hes1,

Vsx2, and Lhx2) and positive regulators of cell cycle progression (various Cyclins) were found to be marked by G9a-dependent H3K9me2 in their promoters, leading to their repression during differentiation. It is not know, however, whether the role of G9a in

RPCs reflects a more general role elsewhere in progenitors of the CNS, or what (if any) are the identities of possible RPC-specific cofactor proteins/ncRNAs that regulate G9a in the CNS. It is into both of these aspects of G9a function in the developing CNS that my thesis sheds particularly interesting insights.

1.4 Aims and objectives

This work focuses on studying protein complexes responsible for controlling developmental gene expression programs via the catalysis and/or recognition of specific histone methylation marks. When I started my graduate work in the winter of 2008, a subset of histone PTMs, HMTs and methyl-lysine binding protein domains were under intense investigation, particularly the structural aspects underlying their substrate specificities. While the composition and function of certain histone methylation-related complexes had already been characterized in yeast and, to a lesser extent, in Drosophila, little was known regarding orthologous complexes and corresponding functions in mammalian cells. In particular, it was largely unknown what human proteins physically associate with histone methylation-related enzymes, how these regulate chromatin activity, and in which physiological context. To address this gap in understanding, my work over the past ~6 years has largely been dedicated toward 2 primary aims:

(1) To better understand the composition of protein complexes and the networks of physical interactions that comprise the histone methylation- and transcription-related machinery responsible for developmental gene expression programs; and

(2) To determine the biological roles of these protein complexes and their individual components within the context of developing biological systems.

Chapter 2 A physical interaction map for histone methylation-related proteins

The work presented in this chapter was generated as part of a close collaboration between the Emili and Greenblatt labs. I received technical assistance from some members of the Greenblatt lab, particularly Guoqing Zhong and Xinghua Guo for AP-MS. The work related to the RPRD proteins constitutes my contribution to a paper published in the journal Transcription (Ni et al., 2011). The work in this chapter related to histone methylation-related complexes (together with the work in Chapter 3) is part of a paper to be published in the near future.

2 A physical interaction map for histone methylation-related proteins 2.1 Abstract

Histone methylation-related complexes are key regulators of gene expression during development, yet their core composition and the identities of tissue-specific cofactors that regulate their functions are incompletely understood. To better characterize the components of histone methylation-related complexes, I applied a systematic lentiviral-based affinity purification-mass spectrometry (AP-MS) approach in human cell culture to a target set of proteins known or predicted to regulate histone methylation and developmental gene expression (listed in Appendix 1). After stringent data processing and filtering, the resulting high-confidence physical interaction network recapitulated many novel (261) and previously annotated (312) protein-protein interactions and curated protein complexes. Among the unexpected histone methylation- and transcription-related protein assemblies found in my proteomic survey were complexes containing previously uncharacterized proteins: (1) RPRD1A, RPRD1B and RPRD2 in association with

RNAPII, and (2) ZNF644, a functionally unannotated zinc-finger-containing protein recently linked to high-grade myopia that specifically associated with G9a/GLP. I found that RPRD1A, RPRD1B, and RPRD2 are components of an RNAPII-containing protein complex that contains the putative CTD phosphatase RPAP2 and the transcriptional co- factors GRINL1A and RECQL5. In addition, I found that ZNF644 binds to G9a/GLP via an atypical ZF and thereby co-regulates global H3K9me2 levels. Collectively, this chapter of my thesis work describes a high quality PPI reference map for the human

histone methylation-related protein machinery and highlights important novel regulators of transcription.

2.2 Introduction

This chapter focuses on my work related to a proteomic survey of histone methylation-related protein interactions and protein complexes using high performance

AP-MS. Mass spectrometry-based proteomics has become a proven and successful tool for the identification of multi-protein complexes (Gingras et al., 2007) and an increasingly important method for the systematic mapping of intracellular protein machinery. AP-MS has been broadly applied to interactome mapping in microbes such as budding yeast (Ho et al., 2002; Gavin et al., 2006; Gavin et al., 2002; Krogan et al., 2006) and, to a more limited extent, in cultured cell lines (Hutchins et al., 2010; Ewing et al.,

2007; Bouwmeester et al., 2004; Jeronimo et al., 2007; Sowa et al., 2009; Guruharsha et al., 2011). For my studies, I made use of a newly introduced lentiviral-based system, termed MAPLE (Mammalian Affinity Purification using Lentiviral Expression), developed in collaboration by the Moffat, Greenblatt and Emili labs at the Donnelly

Centre (Ni et al., 2011; Mak et al., 2010). In this system, outline schematically in Figure

2.1, a sequence verified open reading frame (ORF) of interest is tagged at either the N- terminus or C-terminus using a versatile triple affinity purification (“VA” or “VAP”) epitope tag cassette. Lentivirus expressing each fusion protein under the control of a constitutive CMV promoter is then produced by recombineering and used to infect mammalian cell culture (i.e., HEK293 cells) to generate integrated cell lines that stably express the epitope tagged constructs. These cells are then harvested, lysed, and the resulting protein lysate subject to two-step affinity purification using the 3xFlag and

2xStrep capture beads. Following trypsin digestion, the peptide mixtures are separated using reverse phase chromatography and electrosprayed into a tandem mass spectrometer

(in my case, a high precision Orbitrap-Velos hybrid instrument). The resulting peptide fragmentation mass spectra were searched against a theoretical “human peptidome” using the SEQUEST algorithm and high confidence matches selected using the STATQUEST probabilistic model (Kislinger et al., 2003). Finally, putative co-complex physical interactions were scored using several previously established algorithms, such as

Significance Analysis of INTeractome (SAINT) (Choi et al., 2011), to determine high- confidence interactions. I describe key aspects of the experimental pipeline, noteworthy features of the resulting high quality network, and follow up validation studies below.

Figure 2.1: Workflow outline of the lentiviral-based AP-MS approach used in this study.

2.3 Methods

2.3.1 Plasmid construction

Most ORFs for histone-related proteins were obtained in a format suitable for

Gateway™ based recombineering (pDONR221 or pDONR223 plasmids) from the

Human ORFeome collection (Open Biosystems), the Ultimate ORFeome collection

(Invitrogen), or the PlasmID Collection (Harvard). In a few instances, ORFs were generated using PCR amplification from cDNA derived from HEK293 cells and then cloned into Gateway™ vectors using the BP reaction (Invitrogen, Inc.). Primer sequences used for PCR amplification are found in Table 2.1. The C1263A and H1283A point mutant versions of full-length ZNF644 (note: C1263 and H1283 are putative zinc ion coordinating residues of the C-terminal atypical ZF motif) were generated using the

Phusion Site-Directed Mutagenesis Kit (F-541, Thermo Scientific, Inc.) as per the manufacturer’s protocol using the template pDONR223-ZNF644 (isoform 1, described above). The primers for SDM, listed in Table 2.2, were HPLC purified and 5’ phosphorylated. All ORFs were verified by plasmid DNA sequencing (Bio Basic).

2.3.2 Cell culture and lentivirus production

HEK293 and HEK293T cells were cultured at subconfluence in Dulbecco’s

Modified Eagle Medium (DMEM) with 10% fetal bovine serum (FBS) and antibiotics as previously described (Moffat et al., 2006; Mak et al., 2010). Lentivirus was produced and infected at a multiplicity of < 1 essentially as previously described (Moffat et al., 2006).

Table 2.1: Primers used to clone into Gateway™ isoforms 1 (full-length) and 3 (C- terminal region only) of human ZNF644.

Primer Forward (5’à3’) Reverse (5’à3’)

ZNF644 GGGGACAAGTTTGTACAA GGGGACCACTTTGTACAA (isoform 1) AAAAGCAGGCTCAATGAG GAAAGCTGGGTTCTATGA ATCGTTCTTGCAGCAAG AGCTGCTTCGGCC

ZNF644 GGGGACAAGTTTGTACAA GGGGACCACTTTGTACAA (isoform 3) AAAAGCAGGCTCAATGCT GAAAGCTGGGTTCTATGA GATTCGTCAGAACC AGCTGCTTCGGC

Table 2.2: Primers used for site-directed mutagenesis of putative zinc-ion coordinating residues of ZNF644.

Primer Forward (5’à3’) Reverse (5’à3’)

ZNF644-C1263A CATTCTCCGAGCCAGGTTTTGT TAAACCTCGTCA GCACCATGAGG

ZNF644-H1283A ACTGGATTAAGGCCTTACAACGAC CTTCCTGAACAG ACAAGGGTCCT

2.3.3 Affinity purification and mass spectrometry

AP-MS assays were performed essentially as previously described (Mak et al.,

2010; Ni et al., 2011; Walkey et al., 2012). Briefly, bottom-up shotgun sequencing of tryptic peptide mixtures was performed on an Orbitrap-Velos mass spectrometer

(ThermoFischer Scientific) using collision-activated dissociation (CAD) after separation

by nanoflow reverse phase chromatography. Ionization was achieved using a positive electrospray voltage of +2.5 kV applied to a capillary nanospray ion source (Proxeon).

The instrument was operated in a targeted data acquisition mode, with each high resolution (60,000) full mass scan followed by 10 sequentially acquired data-dependent

MS/MS spectra. A dynamic target exclusion list was enabled to minimize redundant peptide sampling. The RAW files were extracted with the ReAdW program and submitted for database searching using a cluster computer distributed version of

SEQUEST (v2.7) against a modified UniProtKB/Swiss-Prot fasta sequence file bearing additional entries as follows: BSA (P026769); GFP (P42212); TEV (P04517) and

Streptavidin (P22628). The SAINT and compPASS scoring algorithms were obtained and applied as previously described (Choi et al., 2011; Skarra et al., 2011; Sowa et al., 2009).

2.3.4 Immunoprecipitation and western blotting

Cell lysates were incubated overnight at 4°C with 2 µg of antibody followed by incubation with 20 µl Protein G beads (Sigma) for 4 hours. After washing with low salt buffer (10mM Tris-HCl, pH 7.9, 100mM NaCl, 0.1% NP-40), proteins were eluted into protein loading buffer, followed by western blot. The following monoclonal antibodies were used for immunoprecipitation: Flag (Sigma, F1804), Gal4 (Millipore, 06-262), total

RNAPII (N-20, Santa Cruz, SC-899), non-phosphorylated RNAPII CTD (8WG16,

Greenblatt lab), CTD-S2P (3E10, Eick lab), CTD-S5P (3E8, Eick lab), and CTD-S7P

(4E12, Eick lab). Western blot assays were performed as previously described (Mak et al., 2010). Immunoblotting was performed as previously described (Mak et al., 2010).

2.3.5 Lentiviral-based shRNA-mediated mRNA knockdown

shRNA knockdown clones were produced in lentivirus as previously described

(Moffat et al., 2006). An shRNA targeted against firefly luciferase (shLuc) was used as a non-silencing control as previously described (Moffat et al., 2006). shRNA constructs targeted against G9a and ZNF644 were kind gifts from Jason Moffat. Knockdown efficiency was assessed using quantitative PCR or Western blot (see below).

2.3.6 Quantitative PCR

Total RNA extraction and qRT-PCR were performed in technical triplicate as previously described (Mak et al., 2010), with the following modification: standard curves for each primer set was generated by serial dilution of a mixture of equal amounts of each cDNA from the sample set being analyzed on an 7300 Real-Time PCR System (Applied

Biosystems). Data was acquired using the following program: (1) 1 cycle, 95°C, 10min;

(2) 40 cycles, 95°C, 15s; (3) 1 cycle 55°C, 30s. GAPDH and B-actin transcripts were amplified in parallel and used as references for relative quantitation via the ΔΔCt method

(built-in software application).

2.4 Results

2.4.1 A physical interaction map for histone methylation-related proteins

Shortly after beginning my thesis work, a collaborative effort between the Moffat,

Greenblatt and Emili labs led to the development of an effective and efficient AP-MS method for the characterization of protein complexes involved in various aspects of transcriptional regulation (Mak et al., 2010; Ni et al., 2011). I decided to apply this method systematically to components of human histone methylation-related protein

complexes using HEK293 cells, which exhibit many characteristics of immature neurons

(Shaw et al., 2002), that stably express epitope-tagged “bait” proteins known or predicted to regulate histone methylation and/or developmentally regulated gene expression programs. These included confirmed or putative homologs of PcG, COMPASS-like, and

G9a/GLP-containing protein complexes described in Chapter 1 (also see the Appendix

1 for the complete AP-MS dataset). Since over-expression of bait proteins can alter the composition of protein complexes, such as inducing physical interactions that normally do not occur, I used reciprocal AP-MS or by IP-WB to validate select novel interactions.

After eliminating from consideration baits that failed to generate meaningful AP-MS data

(e.g., due to insufficient expression of the bait or impaired cell viability), I assembled a high-confidence dataset comprising 33 successful replicate AP-MS experiments encompassing 25 histone methylation-related and 8 transcription-related bait proteins principally as positive controls.

To assemble the PPI data into binary networks of pairwise interactions, I applied a multipronged approach using recently published “spoke” model-based algorithms, namely CompPASS (Comparative Proteomic Analysis Software Suite) (Sowa et al.,

2009) and SAINT (Statistical Analysis of Interactome) (Choi et al., 2011). In contrast to

“matrix” models, which assign all possible pairwise PPIs between the proteins identified in a given purification (i.e., both bait-prey and prey-prey PPIs), spoke model-based methods only assigns bait-prey interactions. While others have applied matrix-based models to AP-MS datasets (Guruharsha et al., 2011), my dataset contained several bait proteins known to be components of biochemically and functionally distinct complexes.

For example, AP-MS experiments for WDR5 purifies components of the histone

acetylation Ada2a-containing (ATAC) complex (Suganuma et al., 2008), various

COMPASS-related complexes, and other distinct complexes. Likewise, certain subunits of RNAPII also reside in RNAPI and RNAPIII complexes (Ni et al., 2011). After generating probabilistic networks, I opted to apply the SAINT scoring algorithm at a cut- off value of 0.83 to filter the dataset (i.e., to remove background contaminant proteins) as this threshold score accurately recapitulated the known compositions of well-established transcription complexes, such as Mediator (Conaway et al., 2005), RNA polymerase II

(RNAPII) (Ni et al., 2011) and the cleavage and polyadenylation specific factor (CPSF) termination complex (Kaufmann et al., 2004; Takagaki and Manley, 2000) (Figures 2.2 and 2.3). In addition to these positive-control transcription-related complexes, many of the relevant annotated physical interactions previously reported in the public protein- protein interaction repositories BioGRID (Chatr-Aryamontri et al., 2013) and iRefWeb

(Turner et al., 2010) were recapitulated in my filtered PPI dataset (312/428, 72.9%), supporting the reliability of the final interaction network encompassing 573 putative bait- prey co-complex interactions.

To better define complex memberships, I performed two-dimensional hierarchical clustering of the interaction profiles between bait proteins. My results showed aggregation of clusters along the main diagonal, and, as expected, the interaction profiles for protein subunits known to reside in the same complex(es) were characteristically highly correlated (Figure 2.4). Hence, I concluded that my PPI dataset recapitulated well-characterized histone methylation-related and transcription-related protein complexes while enabling the assignment of putative interesting novel components.

A B

Figure 2.2: Computational scoring and filtering approach for the transcription- and histone methylation-related AP-MS dataset used in this study. (A) SAINT and compass scores of annotated (red) and non-annotated (black) protein interactions from the histone methylation- and transcription–related AP-MS dataset. (B) SAINT-based filtering approach used to generate a high-confidence protein interaction network.

Figure 2.3: Physical interaction network for histone methylation- and transcription- related protein complexes. Physical interaction map based on AP-MS data for histone methylation-related complexes. Nodes represent individual proteins identified by LC-

MS/MS. Black edges represent interaction reported in my dataset, and green edges represent previously reported interactions from iRefWeb (Turner et al., 2010) and/or

BioGRID (Chatr-Aryamontri et al., 2013)

HP1

UTX/COMPAS

PRC1/BCORS

PRC2

Mediator Correlation 0 1 RNAPII

Figure 2.4: Hierarchal clustergram of the pair-wise correlation of the interaction profiles of the indicated bait proteins. Bait proteins that are known to reside within the same or similar protein complexes are grouped in colored boxes. Numerical correlation score values are listed in Appendix 1.

2.4.2 RPRD1, RPRD1B, and RPRD2 as novel RNAPII-interacting transcriptional co-factors

Initially intended as a positive control, I performed AP-MS experiments using tagged alleles of the POLRD2 and POLR2E subunits of RNAPII. As expected, I recovered most of the annotated subunits of RNAPII and, in the case of POLR2E, shared subunits of RNAPI and RNAPIII (refer to Appendix 1 for AP-MS dataset). Strikingly, however, I identified three previously uncharacterized proteins co-purifying with both

POLR2D and POLR2E, namely RPRD1A, RPRD1B and RPRD2 (referred to, generally, as the RPRD proteins (Note: RPRD is an acronym for “Regulation of Nuclear Pre-mRNA

Domain-Containing”) with significant supporting mass spectral evidence (Table 2.3). It had been reported that RPRD1A, also called P15RS, regulates certain cell cycle genes

(Liu et al., 2002). Later, it was demonstrated that RPRD1A can attenuate Wnt signaling by disrupting the interaction of beta-catenin and TCF4 (Wu et al., 2010). However, the physical associations of RPRD1A (or RPRD1B and RPRD2) with RNAPII have not been previously reported, and the RPRD proteins were largely functionally uncharacterized when I identified them.

Table 2.3: Summary of select AP-MS data for the POLR2D and POLR2E subunits of RNAPII, RPRD1A and RPRD1B. Numerical values represent averaged spectral counts from duplicate AP-MS experiments. Refer to Appendix 1 for the complete dataset for transcription-related proteins.

Baits Preys POLR2D POLR2E RPRD1A RPRD1B MED1 13.5 19.5 0.0 0.0 MED14 0.0 21.0 0.0 0.0 MED17 8.5 18.0 0.0 0.0 MED19 0.0 4.5 0.0 0.0 MED21 6.0 9.0 0.0 0.0 MED30 0.0 4.0 0.0 0.0 MED4 0.0 9.5 0.0 0.0 MED8 4.5 6.5 0.0 0.0 PHRF1 4.5 0.0 0.0 0.0 POLR2A 534.5 662.5 375.0 320.0 POLR2B 414.5 439.0 262.5 214.5 POLR2C 98.0 145.0 66.5 64.0 POLR2D 85.0 20.5 16.5 25.5 POLR2E 85.0 97.5 35.0 31.0 POLR2G 205.0 61.0 30.0 30.5 POLR2H 62.5 197.0 36.0 36.0 POLR2I 48.0 59.5 16.5 24.0 POLR2J 29.5 21.5 12.5 23.5 POLR2L 0.0 9.5 0.0 2.0 POLR2M 24.0 28.0 16.0 19.5 RECQL5 33.0 0.0 25.0 40.5 RPAP2 28.0 0.0 28.5 27.0 RPRD1A 94.0 82.0 627.5 116.5 RPRD1B 68.5 89.5 61.5 223.5 RPRD2 86.5 113.0 17.0 118.0 SCAF4 6.0 12.5 0 4.0 SUPT6H 10.5 5.5 10.5 10.5

The RPRD proteins are evolutionary conserved and represent 3 of the 8 CID- containing proteins annotated in humans (Figure 2.6A). The CIDs of the RPRD proteins appear to be most closely related to the yeast protein Rtt103 (which lacks an obvious mammalian orthologue), the CID of which been structurally characterized in complex with a CTD derived S2P heptapeptide (Lunde et al., 2010). As seen in Figure 2.5A,

RPRD1A and RPRD1B are largely composed of the same protein domains and share a high degree of sequence similarity throughout the proteins. RPRD2, however, is much larger and contains compositionally biased serine- and proline-rich regions, the latter of regions of which are reported to be phosphorylated during M-phase (Olsen et al., 2010).

The proline-rich region of RPRD2 contains multiple repeats of the consensus sequence

PPPPP[D/E]H, which has been shown to be a peptide recognition sequence for various

WW domains, including the TF and putative tumor suppressor WWOX (Ingham et al.,

2005).

B C

WB: Flag (VA-RPRD1A) Flag (VA-RPRD1B) POLR2D

* * * * * * * * * * *

Figure 2.5: The RPRD proteins specifically bind phosphorylated RNAPII. (A) Protein architecture of RPRD1A, RPRD1B and RPRD2 as found in the UniProt database. (B)

Averaged spectral counts from duplicate AP-MS experiments for VA-tagged RPRD1A or

RPRD1B from HEK293 cells. Asterisks indicate core subunits of RNAPII. (C) IP-WB experiments from HEK293 cells using the indicated IP and WB antibodies. Note that

POLR2D blots are from WT HEK293 cells, whereas Flag-RPRD1A and 8WG16 binds unphosphorylated CTD repeats, N-20 binds the N-terminal portion of POLR2A (i.e., total

RNAPII), 3E10, 3E8 and 4E12 bind S2P, S5P and S7P CTD repeats, respectively.

To confirm the interactions between the RPRD proteins and RNAPII, I performed reciprocal AP-MS experiments using lentivirus expressing tagged RPRD1A and

RPRD1B as baits. As predicted, proteins identified as stably co-purifying with RPRD1A and RPRD1B from HEK293 cells included almost all core RNAPII subunits, but I also detected GRINL1A, which creates a Mediator requirement for transcriptional activation in vitro reconstitution experiments (Hu et al., 2006), the helicase RECQL5, which inhibits transcription initiation by RNAPII in vitro (Aygün et al., 2009; Kanagaraj et al.,

2010), and the putative CTD phosphatase RPAP2 (Mosley et al., 2009), which has been proposed to target the S5 residue (Figure 2.5B). RPRD1A and RPRD1B likewise co- purified with each other and with RPRD2 (Figure 2.5B). Conversely, specific subunits of

RNAPI, RNAPIII, and the Mediator complex, which binds RNAPII with an unphosphorylated CTD (Shi et al., 2011) were not detected. RPRD proteins were also detected in reciprocal AP-MS experiments for GRINL1A, but not when Mediator subunits were tagged and purified (Appendix 1). Additional results from immunoprecipitation and Western blotting (IP-WB) experiments confirmed the interactions of endogenous RPRD1A, RPRD1B and GRINL1A with untagged RPAP2

(Ni et al., 2011). Collectively, these data suggest the formation of a novel RNAPII- associated protein complex containing the RPRD proteins, GRINL1A, RPAP2 and

RECQL5.

The RPRD proteins contain CIDs, which generally bind phosphorylated CTD peptides, leading me to hypothesize that the RPRD proteins might likewise recognize phosphorylated forms of RNAPII. To test this possibility, IP-WB experiments were performed using different using monoclonal antibodies that recognize total RNAPII (N-

20) or specific CTD phosphorylation marks: 4E12 (unphosphorylated), 3E10 (S2P), 3E8

(S5P), 4E12 (S7P) (Chapman et al., 2007). Indeed, the immunoprecipitation experiments demonstrated that RPRD1A and RPRD1B both selectively co-immunoprecipitated with all three phosphorylated CTD variants, but not with unmodified RNAPII (Figure 2.5C).

These results demonstrate that RPRD1A and RPRD1B predominately bind to RNAPII with a phosphorylated CTD, the significance of which is further explored in the

Discussion section.

2.4.3 BOD1L as a novel component of a COMPASS-like complex

In addition to the physical interactions of RNAPII with the RPRD proteins described above, my interaction network revealed several putatively novel physical interactions among histone methylation-related proteins. For example, AP-MS experiments using multiple individual subunits of histone human COMPASS-like complexes identified several annotated components as well as the uncharacterized protein

BOD1L (Appendix 1). I found that BOD1L specifically co-purified with the core

COMPASS subunits ASH2L and WDR5, but not with the MLL2/MLL3-specific histone demethylase subunit, KDM6A/UTX. It should be noted that additional COMPASS-like subunits were attempted (i.e., DPY30, RBBP5) but were not processed further due to difficulties detecting bait expression in culture (data not shown). Using IP-WB, Dr. Edyta

Marcon, a postdoctoral researcher in the Greenblatt lab, validated the interaction between the endogenous BOD1L protein and epitope-tagged WDR5 (Figure 2.6A). Similar findings regarding the association of BOD1L with a specific type of COMPASS-like complex were recently reported (van et al., 2013), yet much remains unknown as to how

BOD1L might regulate the activity of COMPASS-like complexes.

2.4.4 A novel SFMBT1-L3MBTL3 heteromeric co-repressor complex

In addition to COMPASS-like complexes, my interaction network indicated several putatively novel interactions for various PcG-like proteins, such as SFMBT1 and

L3MBTL3, which are the mammalian versions of the Drosophila dSfmbt and L(3)mbt, respectively. It is now known that SFMBT1 and L3MBTL3 are able to bind histone methylation marks in vitro via their MBT domains (Nady et al., 2012). However, in contrast to the highly studied core components of PRC1 and PRC2 protein complexes,

Sfmbt resides in a less well-known Pho-repressive complex (PhoRC) that appears to lack a direct mammalian counterpart (Wu et al., 2007; Cai et al., 2007). However, my AP-MS experiments for SFMBT1 and L3MBTL3 revealed a novel protein complex comprising these two methyl-lysine “readers” in stable association with the histone demethylase

KDM1A/LSD1, the SAM domain-containing protein SAMD1, and the Co-REST repressors RCOR1 and RCOR3 (Appendix 1, Figure 2.6B). KDM1A/LSD1 has been reported to target both activating (H3K4) and repressive (H3K9) methylation marks, but when in complex with RCOR1/RCOR3, LSD1/KDM1A appears to preferentially function as a transcriptional repressor by targeting H3K4 for demethylation (Shi et al.,

2005; Lee et al., 2005). Consistent with my findings, it was recently demonstrated that a compositionally similar SFMBT1-L3MBTL3-KDM1A/LSD1 complex is formed in germ cells during spermatogenesis, where the assembly (or at least its subunits SFMBT1 and

KDM1A/LSD1) down-regulates the expression of canonical histone genes (Zhang et al.,

2013).

A IP

IgG

Irrelevant Irrelevant 10% Input Flag

L3MBTL3 LSD1 WB:

Flag (VA-SFMBT1)

B IP

IgG

Flag 10% Input Irrelevant Irrelevant BOD1L MLL2

WB:

Flag (VA-WDR5)

Figure 2.6. Validating interactions derived from my AP-MS dataset by IP-WB. (A) IP-

WB experiment from HEK293 cells stably expressing VA-tagged PcG-like protein

SFMBT1 using the indicated antibodies for either IP or WB. (B) IP-WB experiment from

HEK293 cells stably expressing VA-tagged WDR5 using the indicated antibodies for either IP or WB. Data provided by Dr. Edyta Marcon (Greenblatt lab).

2.4.5 ZNF644 as a novel G9a/GLP-interacting protein

Most notable of all my purifications, I found that VA-tagged G9a reproducibly co-purified with not only its known heterodimerization partner GLP (Tachibana et al.,

2005; Tachibana, 2005), but also two large vertebrate-specific C2H2-like ZF proteins,

WIZ and ZNF644 (Figure 2.7, Appendix 1). While WIZ had already been reported to interact with G9a and GLP in ES cells, where it stabilized G9a protein levels (and therefore regulated its activity) and bridged the HMT to the CtBP co-repressor complex

(Ueda et al., 2006), ZNF644 was entirely novel. To validate the physical association of

G9a with ZNF644, I performed additional AP-MS experiments using HEK293 cells expressing GFP-tagged ZNF644 under the control of a doxycycline-inducible promoter

(See Flp-In TRex section in the Methods section). Indeed, as expected, GFP-tagged

ZNF644 reproducibly co-purified both G9a and GLP (Figure 2.8A), confirming the stable physical association of ZNF644 with the HMTs. In addition, ZNF644 co-purified with WIZ (Figure 2.8A), suggesting that ZNF644 and WIZ may also reside within the same protein complex. These data demonstrate that ZNF644 is a novel interacting partner for the H3K9 HMTs G9a and GLP.

Figure 2.7: G9a co-purified two large C2H2-like ZF-containing proteins, WIZ and

ZNF644. (Top) Protein domain architecture of WIZ and ZNF644, which were identified in AP-MS experiments for VA-tagged G9a from HEK293 cells. Black boxes indicate the positioning of the C2H2-like ZF motifs within ZNF644 and WIZ. (Bottom) The atypical

C2H2-like ZF motifs of WIZ and ZNF644, their amino acid sequence with emphasis on their similarity, and putative zinc ion coordinating residues are indicated.

A Baits

WT ZF8 - - C1263A H1283A

- - GFP ZNF644 ZNF644 - - ZNF644 ZNF644 - - GFP GFP

Preys GFP GFP

GFP 160.0 19.0 20.5 27.0 46.0 ZNF644 0.0 136.0 74.0 87.5 38.0 G9a 0.0 33.0 0.0 0.0 218.0 GLP 0.0 15.0 0.0 0.0 147.0 WIZ 0.0 11.5 0.0 0.0 95.0

-WT -C1263A -H1283A ZF8 ZF8 - -ZF8 - Input (10%)GST GST GST GST

His (G9a SET)

Figure 2.8: The atypical C2H2-like ZF motif of ZNF644 directly binds to the

G9a/GLP SET domain. (A) Averaged spectral counts for the indicated prey proteins from duplicate AP-MS experiments from HEK293 cells stably expressing GFP-tagged

WT, C1263A, H1283A or ZF8 versions of ZNF644. (B) GST pull-down assay using recombinant GST-tagged ZF8 region of WT ZNF644, the C1263A or the H1283A point mutants and the His-tagged G9a SET domain.

2.4.6 ZNF644 binds G9a/GLP via an atypical C2H2-like ZF motif

To characterize the functional importance of a physical interaction, it is helpful to understand which protein domains or binding motifs (contact residues) mediate binding.

It had been previously demonstrated in ES cells that an “atypical” C2H2-like ZF motif situated in the C-terminal region of WIZ is sufficient to co-purify with G9a and GLP

(Ueda et al., 2006). Compared to the consensus sequence of C2H2-like ZF motifs where the zinc ion coordinating residues are separated by a 12-amino acid linker (C-[X2]-C-

[X12]-H-[X3-5]-H), the C-terminal C2H2-like ZF motif of WIZ is “atypical” in that the putative zinc ion coordinating residues are separated by an extended 16-amino acid linker region (C-[X2]-C-[X16]-H-[X3-5]-H). As I investigated the protein sequence of WIZ and

ZNF644, it became apparent that ZNF644 also contained an atypical ZF motif in its C- terminus (Figure 2.7), thus pointing to the possibility that this atypical C2H2-like ZF motif may likewise mediate the G9a-ZNF644 physical interaction.

To determine if the atypical ZF motif of ZNF644 was indeed important for the physical association of ZNF644 with G9a/GLP, I generated alanine point mutations in the putative zinc ion coordinating residues in the atypical ZF motif of ZNF644 (C1263A and

H1283A). I then generated HEK293 cells that expressed integrated copies of GFP-tagged

ZNF644-C1263A or GFP-ZNF644-H1283A and induced their expression by doxycycline treatment. While both proteins were expressed to near wild-type (WT) levels, as evident by the spectral counts for the GFP tag (Figure 2.8A), I found that neither the C1263A nor the H1283A mutant version of ZNF644 was able to co-purify G9a or GLP and that both mutant versions likewise failed to co-purify WIZ (Figure 2.8A). These data suggest that the atypical C2H2-like ZF motif of ZNF644 likely requires the formation of a C2H2-like

ZF-like secondary structure and mediates the physical association of ZNF644 with G9a,

GLP and WIZ. Furthermore, AP-MS experiments using a shorter isoform of ZNF644

(isoform 3) that consists of only the C-terminal region encompassing the atypical ZF motif (ZF8; see Methods section) efficiently co-purified G9a, GLP and WIZ (Figure

2.8A). Collectively, these data demonstrate that the atypical C2H2-like ZF motif of

ZNF644 is sufficient for its stable physical association with the HMTs G9a and GLP as well as WIZ and that this interaction likely requires the formation of a C2H2-like ZF secondary structure.

To determine whether the atypical ZF motif of ZNF644 directly binds to G9a, I purified a recombinant form of the region of ZNF644 encompassing the atypical ZF motif region (residues 1213-1314) as a GST fusion protein in E. coli and performed GST pull-down assays with recombinant purified His-tagged G9a catalytic SET domain

(residues 913-1193). The G9a SET domain forms a stable homodimer in vitro that is commonly used as a proxy for the G9a-GLP SET heterodimer (Wu et al., 2010). It has also been shown that the SET domains of G9a and GLP are required for association with the atypical C2H2-like ZF region of WIZ (Ueda et al., 2006), although whether this affects G9a catalytic activity or substrate binding is currently unknown. Indeed, the atypical C2H2-like ZF motif region of ZNF644 stably co-purified with the G9a SET domain, and this binding required a C2H2 ZF-like secondary structure because mutating the putative zinc ion-coordinating residues (C1263A or H1283A) disrupted the interaction (Figure 2.8B). This data suggests that the atypical C2H2-like ZF motif of

ZNF644 directly binds the dimerized SET domain of G9a.

2.4.7 ZNF644 regulates global levels of H3K9me2

Because ZNF644 physically associates with G9a and GLP, and because

G9a/GLP primarily determine H3K9me2, I sought to determine whether ZNF644 likewise regulates H3K9me2 levels. To this end, I created HEK293 cell lines stably expressing short hairpin (sh)RNA targeted against G9a or ZNF644. Indeed, shRNA reducing the expression of G9a or ZNF644 expression both led to reduced levels of

H3K9me2, but the latter did not indirectly mediate this effect by destabilizing either G9a or interfere with H3K9me3 levels (Figure 2.9A). There were no significant changes in monomethylated H3K56, a recently reported G9a histone substrate (Yu et al., 2012), in either G9a or ZNF644 knockdown lines (data not shown). Collectively, these data suggested that ZNF644 directly binds the G9a SET domain homodimer (and likely the

G9a/GLP SET domain heterodimer) and serves as a specific co-regulator of the

H3K9me2 mark.

A B

100%

75% * * 50%

25%

2 1 - - PercentZNF644 transcript remaining relative control shRNA remainingrelative control shRNA

shControl shZNF644 shZNF644

Figure 2.9: ZNF644 is a co-regulator of H3K9me2. (A) qPCR experiments monitoring the levels of ZNF644 transcripts from HEK293 cells stably expressing shRNA targeted against ZNF644 or against luciferase (non-silencing control). Experiments were performed in biological and technical triplicate. Asterisks indicate p-value < 0.05,

Student’s T-test. (B) Western blots performed using the indicated antibodies in HEK293 cell lines stably expressing shRNA targeted against the indicated transcripts

2.5 Discussion

2.5.1 Physical interaction map of the histone methylation system

Recent developments in AP-MS provided me with an unbiased, semi-quantitative and comprehensive means of performing interaction studies on HMTs of interest. Using an innovative lentiviral-based AP-MS approach, I created a physical interaction map encompassing many components of histone methylation- and transcription-related protein complexes with an emphasis on developmental regulators of gene expression. The dataset recapitulated annotated transcription-related protein complexes, such as Mediator,

RNAPII and CSPF complexes, and many other reported protein-protein interactions

(Figures 2.3 and 2.4 and Appendix 1). In addition, I successfully recapitulated the composition of known histone methylation-related complexes, including the core components of PRC2, PRC1, COMPASS-like and G9a/GLP-containing complexes.

Strikingly, however, I identified novel associated complex subunits that motivated further mechanistic investigations into the physiological contexts by which these interactions modulate gene regulation.

2.5.2 The RPRD proteins bind phosphorylated CTD and possibly regulate transcription elongation and/or termination

Perhaps most surprisingly, considering all the research that has been performed on

RNAPII, I identified three evolutionary conserved, but previously uncharacterized, proteins, RPRD1A, RPRD1B and RPRD2, in association with RNAPII molecules containing all three types (S2, S5, and S7) of CTD phospho-epitopes, but not unphosphorylated RNAPII.

Recent unpublished work from my collaborators in the Greenblatt lab demonstrates that the RPRD proteins preferentially bind CTD peptides phosphorylated at

S2 (manuscript in submission), a mark that is most commonly associated with transcriptional elongation and termination. As a result, I propose the following working model for how the RPRD protein may play a role in transcription elongation and termination by RNAPII. First, as RNAPII escapes the promoter regions (which are typically marked by CTD phosphorylation at S5) and enters into productive elongation, the CDK9 subunit of the enzyme P-TEFb (as well as CDK12 and CDK13 (Bartkowiak et al., 2010)) phosphorylates the CTD at S2. The RPRD proteins (via their CIDs) recognize

this phosphorylation mark, resulting in the co-recruitment of other co-complexed proteins

(RECQL5, GRINL1A, and RPAP2). As a putative phosphatase for the S5 phosphorylation mark, RPAP2 may aid in removing the S5 phosphorylation mark from the CTD and thus facilitate the recruitment of other protein complexes that facilitate transcription elongation and termination. The Greenblatt and Emili labs are currently performing more detailed characterization of these proteins to determine more precisely how they regulate transcription by RNAPII (i.e., which genes are targeted and where within a gene body the complex resides).

2.5.3 High-grade myopia protein ZNF644 physically interacts with G9a and GLP

Among the novel physical interactions I identified was the uncharacterized vertebrate-specific C2H2 ZF protein ZNF644, which was linked to high-grade myopia

(Shi et al., 2011; Tran-Viet et al., 2012), with the HMT G9a. Like WIZ, ZNF644 contains an atypical C2H2-like ZF motif (Figure 2.7) that is sufficient for physical interaction with G9a and GLP (Figure 2.8A) and directly binds SET domain homodimer (Figure

2.8B) and likely depends on the formation of a ZF-like secondary structure (Figure 2.8A-

B). This atypical C2H2-lke ZF motif appears to be unique to WIZ and ZNF644, as I was unable to identify any other metazoan (Drosophila, zebrafish, mammalian genomes) proteins that contained motifs of any significance sequence similarity. In addition, despite the ~50 SET domain-containing proteins present in the human genome, the binding of the atypical C2H2-like ZF motif of ZNF644 to the G9a SET domain appeared to be highly specific, as ZNF644 did not co-purify any of the many other SET domain-containing proteins expressed in HEK293 cells. I also found that ZNF644 specifically co-regulated

H3K9me2 levels in human cell culture, but not other histone methylation marks, such as

H3K9me3 (Figure 2.9 and data not shown), suggesting specific function related to gene silencing in euchromatic regions. However, it is not obvious how ZNF644 might impart target gene specificity for G9a/H3K9me2, as its arrangement of C2H2-like ZF motifs does not reflect the typical DNA binding variety. Along these lines, in vitro protein binding microarray (PBM) assays using the C2H2-like ZF clusters of ZNF644 did not show sequence-specific DNA binding activity (data not shown). One possibility is that

ZNF644 might act as an adaptor scaffold protein that bridges the HMTs G9a and GLP to other repressor proteins with more direct targeting capabilities. Indeed, in ES cells, WIZ has been reported to bridge G9a and GLP to the CtBP co-repressor complex in ES cells .

Collectively, these data demonstrate that ZNF644 is a novel G9a interacting protein and a specific co-regulator of the H3K9me2 mark. However, the physiological context in which this physical association might exert its function is not clear from these assays, which provided the motivation for my work as described in Chapter 3.

Chapter 3 G9a and ZNF644 physically associate to suppress progenitor gene expression and cell cycle progression in the retina

This chapter contains data from a collaborative effort between the Emili and Tropepe labs. I initially conceived using zebrafish as a model system for my thesis and received critical insights from Profs. Vincent Tropepe and Andrew Emili regarding experimental design and data interpretation. For these follow up studies, I worked closely with (now Dr.) Loksum Wong, who shared her time, talents, and expertise in zebrafish neural development. It is anticipated that the work presented in this chapter will be published, together with the data in Chapter 2, in the near future (manuscript in submission).

3 G9a and ZNF644 physically associate in neuronal stem/progenitor cells to suppress progenitor gene expression and cell cycle progression in the developing retina 3.1 Abstract

Given my finding of a novel G9a-ZNF644 physical interaction, the link of

ZNF644 to high myopia, and the importance of epigenetic silencing during cell fate specification, I sought to better understand the physiological context(s) in which this newly discovered complex functioned during development. To this end, I turned to a zebrafish embryo model system to characterize the functions of two putative ZNF644 orthologues (27.1% and 19.6% protein sequence identity), termed Znf644a and Znf644b, which I found to be specifically expressed in retinal and midbrain progenitor cells. I determined that znf644a and znf644b are both required for the suppression of certain multipotency and cell cycle-related genes (i.e., vsx2 and ccnd1) as well as H3K9me2 levels in their promoter regions. Upon further exploration of the znf644a and znf644b morphant phenotypes, I observed unexpected distinctions between the two retinal phenotypes – that is, retinal cells in znf644b morphant embryos maintained a prolonged proliferative RPC identity, resulting in the delayed formation of differentiated neurons, whereas znf644a morphant retinal cells, as a population, withdraw from the cell cycle and differentiate into various types of retinal neurons, albeit with impaired cell viability.

Importantly, I used “genetic cooperativity” as well as complementation assays to provide evidence that znf644a and znf644b function cooperatively with g9a and that they are critically dependent on their ability to physically bind G9a. Moreover, I found evidence that the functions of znf644a and znf644b are conserved in mammalian ZNF644 and that

the retinal defects of znf644a and znf644b morphants are recapitulated elsewhere in zebrafish midbrain progenitor cells. Collectively, these findings demonstrate that a G9a-

ZNF644 complex mediates multiple important regulatory functions during neural progenitor cell differentiation.

3.2 Introduction

During development, proliferating progenitor cells withdraw from the cell cycle and generate post-mitotic progeny of specialized function. The correct timing and execution of such cell fate transitions involve dynamic modulation of chromatin modification and gene expression by HMTs and ensure proper tissue formation and structure. However, the composition, roles and regulation of these HMT complexes in determining cell fate decisions in vivo are poorly understood.

In the zebrafish embryonic retina, RPCs are initially uniformly proliferative by virtue of the expression of a complement of master TFs that repress the expression of genes that negatively regulate cell cycle progression (i.e., p27kip1, p57kip2) and restrict lineage potential (pro-neural TFs). Shortly after the 24 hours post-fertilization (hpf) time point, RPCs begin to undergo unidirectional changes in their competence, withdraw from the cell cycle and produce differentiated progeny of specialized function. During this process, the expression of master multipotency-related TFs (i.e., Vsx2) as well as positive regulators of cell cycle progression (i.e., CyclinD1) are silenced, and the expression of

TFs that guide lineage specification become expressed. By 48 hpf, various types of differentiated cell types begin to form, which are identified based on the expression of cell type-specific protein markers. As described in the Chapter 1, G9a is now known to play a critical role in mouse RPCs, where it suppresses the expression of certain RPC-

specific TFs (i.e., Vsx2) as well as various Cyclins (i.e., CyclinD1) (Katoh et al., 2012).

These findings were unknown at the time began my zebrafish work, but proved to provide valuable insights that enabled me to pinpoint specific RPC-specific genes regulated by the G9a-ZNF644 physical interaction.

This chapter is a follow-up to certain findings from Chapter 2. As described earlier, I identified the functionally uncharacterized vertebrate-specific C2H2 zinc finger protein ZNF644, which is putatively causally linked to high-grade (degenerative) myopia

(Shi et al., 2011; Tran-Viet et al., 2012), as a novel G9a/GLP interacting protein and co- regulator of histone methylation. In close collaboration with Dr. Vincent Tropepe and his graduate student Loksum Wong, I found that the corresponding znf644 paralogues in zebrafish are specifically expressed in retinal and midbrain progenitor cells where they function cooperatively during early embryonic development in association with G9a. In addition, I found that these two paralogues play separable, yet related epigenetic roles to ensure the proper coordinated suppression of progenitor identity and cell cycle progression during differentiation. Importantly, these phenotypic abnormalities observed in znf644a and znf644b were efficiently rescued by co-injecting mRNA encoding human

ZNF644, suggesting conserved function across species. These findings reveal that suppression of a proliferative neural progenitor identity involves independent, yet harmonized suppressive functions mediated by an otherwise evolutionarily conserved

G9a-ZNF644 physical association.

3.3 Methods

3.3.1 Zebrafish husbandry

Adult zebrafish (Danio rerio) were maintained at 28°C on a 14-hour light/10-hour dark cycle and housed in an automated re-circulating system (Aquaneering). Embryos were staged as previously described (Kimmel et al., 1995) and reared according to standard procedures (Westerfield, 2000). The wild-type strain used was AB (Zebrafish

International Resource Center). The transgenic strains Tg(HuC:Kaede) [also known as

rw0130a zc7 Tg(elav3:Kaede) ] and Tg(isl2b:GFP) were kind gifts from Dr. Hitoshi

Okamoto (RIKEN Brain Science Institute) and Dr. Chi-Bin Chien (University of Utah), respectively.

3.3.2 Morpholino injections

Commercial antisense morpholinos (MOs) were obtained from Gene Tools, LLC

(Philomath, OR). Znf644a-MO and znf644b-MO were designed to target the splice junction at exon2-intron2, and the working concentrations were determined by a series of dosage-response experiments. Mismatch control MOs (mMOs) were also synthesized for znf644a-MO and znf644b-MO. G9a-MO and p53-MO were as previously described

(Prykhozhij, 2010). A control mismatch MO for g9a-MO was used as previously described (Rai et al., 2010). Morpholino sequences are listed in Table 3.1. Unless otherwise noted, the yolks of 1- to 2-cell stage embryos were injected with empirically optimized amounts of znf644a-MO (30 ng), znf644b-MO (20 ng), g9a-MO (3 ng), and/or p53-MO (2 ng).

3.3.3 Quantitative RT-PCR

Total RNA was extracted from embryos (n=20-40) with Trizol Reagent

(Invitrogen). First strand cDNAs were reverse transcribed from oligo(dT)12-18 primed total RNA (DNase treated) using SuperScript III (Invitrogen). Each 25µl reaction consisted of 1x PCR buffer (1.5mM MgCl2, 0.2mM dNTPs), 0.4uM each of forward and reverse primers, 0.5U Platinum Taq DNA Polymerase, and diluted cDNA template

(1:100). PCR conditions were as follows: 95°C for 5 minutes, followed by 35-40 cycles of 95°C for 30 seconds, 52-58°C for 30 seconds, and 72°C for 30 seconds. Annealing temperatures and cycle number for each primer pair were determined using gradient

PCR. PCR products were resolved on a 1% agarose gel, stained using ethidium bromide, and imaged using a Gel Doc XR+ System and Quantity One Software (Bio-Rad,

Hercules, CA). Primer sequences, listed in Table 3.2, were designed to measure the levels of unspliced nascent RNA and mature spliced mRNA.

Table 3.1. Antisense morpholino sequences used for loss-of-function investigations.

Morpholino Sequence (5’à3’)

znf644a-MO GTGAGCAATAATCACCTTTTCTGAT znf644a-mMO GTGtGCAtAATCtCCTTaTCaGAT

znf644b-MO ATTAAAATTGTCACCTGTTTTGACT znf644b-mMO ATTtAAtTTGTCAgCTGTaTTcACT

g9a-MO GACACACACTGACCTGCAGATGATC

g9a-mMO CCTCTTACCTCAGTTACAATTTATA

p53-MO GCGCCATTGCTTTGCAAGAATTG

Table 3.2. Primer sequences for the indicated unspliced or spliced transcripts.

Target transcript Forward (5’à3’) Reverse (5’à3’)

Spliced/unspliced TCATGGCAGATAAGCCAGAGT TGCCGTCACAAGTGGAGTAG znf644a

Spliced znf644b AAGTTGGTGTAAGTCTAAGCCAGAA GAGCCTTTATTAATCTCTAACCTTTTT

Unspliced znf644b TGAACCGAGCTCAGATGTTG GTCCGTTACCCAGCCTAACA

Spliced g9a AAGTTGGTGTAAGTCTAAGCCAGAA GAGCCTTTATTAATCTCTAACCTTTTT

Gapdh (control) TGCGTTCGTCTCTGTAGATGT GCCTGTGGAGTGACACTGA

3.3.4 Whole-mount in situ hybridization

Embryos were treated with 0.003% of 1-phenyl-2-thiourea (Sigma), which inhibits melanogenesis and thus improves transparency, fixed in 4% paraformaldehyde, and kept in methanol before performing whole-mount in situ hybridization. Samples were treated with proteinase K and hybridized with ~100ng antisense DIG/FITC-labeled RNA probes overnight at 65°C. Excess probes were washed off the next day and embryos were incubated with anti-DIG/FITC antibodies (1:4000, Roche) at 4°C overnight. Color reactions were performed by mixing in nitro-blue tetrazolium and 5-bromo-4-chloro-3’- indolyphosphate (NBT-BCIP, Roche) as substrates to the samples. Images of stained embryos were captured with a Leica MZ16F dissecting microscope with a QIMAGING digital camera and OpenLab software. The following antisense RNA probes were used:

islet1 (Inoue et al., 1994), lef1 (Dorsky et al., 1999), otx2 (Mercier et al., 1995), shh

(Krauss, Concordet and Ingham, 1993) (kind gifts from Dr. Ashley Bruce, University of

Toronto); neurod, pax6a, vsx2 (Thermo Scientific) and p57kip2 (cb961) (Zebrafish

International Resource Center, Oregon).

3.3.5 Histology

Embryos were fixed in 4% paraformaldehyde and rinsed in phosphate-buffered saline solution. For semi-thin sectioning, embryos were first dehydrated using increasing concentrations of ethanol, followed by embedding with increasing concentrations of

Spurr’s resin in ethanol. Embryos were then left to polymerize at 65°C in 100% Spurr’s resin. Semithin coronal sections (approximately 1µm thick) were cut with a glass knife using an ultramicrotome and dried onto glass slides. Counterstaining with toluidine blue to visualize zebrafish morphology followed this procedure. Whole-mount in situ hybridization embryos in 100% glycerol were washed with PBT and followed by the same embedding and sectioning steps as above. Sections were 1.5 µm thick without counterstaining to maximize visualization for the NBT-BCIP precipitate. For cryosectioning, 24-72 hours-post-fertilization (hpf) embryos from each group were fixed with 4% paraformaldehyde overnight at 4°C and washed with a sucrose series (from 5% to 30% sucrose in PBS) for cryoprotection (except for PCNA labeled embryos, which were fixed in 37% formaldehyde:95% ethanol (3:7 ratio) solution). Samples were left in

30% sucrose:optimum cutting temperature compound (O.C.T.) (2:1 ratio) at -20°C before cutting into 14 µm /20µm sections using a cryostat.

3.3.6 Retinal cross-sectional area measurement

Five plastic sections with similar focal plane were chosen to represent each embryo, and images were taken on a Leica DM4500B compound microscope with a

QIMAGING digital camera and OpenLab software. The areas of interest were measured using the program ImageJ. Results represent the average obtained from at least 5 embryos from each group. Statistical analyses between injected and un-injected groups were performed using Student’s t-test. Differences were regarded as significant for p<0.05.

3.3.7 Ectopic gene expression

RNA extracted from whole embryos at 24 hpf was used to prepare cDNA as described above. ORFs for znf644a and znf644b were PCR amplified using the primers listed in Table 3.3. PCR fragments were cloned into pCS2+. The ORFs for wild-type human ZNF644 as well as the C1263A and H1283A point mutant versions were PCR amplified from the pDONR223 plasmid cells into the pCS2+ vector using the same cloning strategy. Templates were linearized with SacII and transcribed in vitro using the mMESSAGE and mMACHINE T7 kit (Ambion). RNA or RNA + MO were injected into the yolks of 1- to 2-cells stage embryos at the concentrations indicated in the text below.

Table 3.3. Primer sequences used to amplify ORFs for znf644a and znf644b for subcloning into the pCS2+ vector.

ORF Forward (5’à3’) Reverse (5’à3’)

znf644a GTCAATCGATATGGAGGACGAAAAAAAAAGG GTCACTCGAGTTAGGAGACCCCCTCAGG (Zebrafish)

znf644b GCTAATCGATATGTCTGCTTTGAAGGAAAGTGC GCTACTCGAGTCATGAGGATGTTTGCATCAC (Zebrafish)

ZNF644 GCTAATCGATATGAGATCGTTCTTGCAGCAAG GCTACTCGAGCTATGAAGCTGCTTCGGCC (Human)

3.3.8 Whole-mount antibody staining

Ten embryos were fixed in 4% paraformaldehyde at 4°C overnight and washed with PBS. Samples were incubated with block solution

(PBS+1%BSA+1%DMSO+0.8%TritonX-100) for an hour an room temperature and overnight in antibody (1:20) at 4°C. Embryos were then washed with PBS+Triton (x%?) and incubated in goat anti-mouse HRP antibody (1:500) overnight. Peroxidase was detected with 3-3’-diaminobenzidine (DAB) and 3% hydrogen peroxide in the dark.

Images of the stained embryos were taken from both a Leica MZ16F dissecting microscope and a Leica DM4500B compound microscope.

3.3.9 Immunohistochemistry

Cryosections were re-hydrated with 1x PBS and blocked for 2 hours in 0.2%

TritonX-100 + 2% goat serum in PBS at room temperature. Primary antibody in block

solution was applied on sections overnight at 4°C. Slides were washed with PBS + 0.1%

Tween-20 and incubated with secondary antibody for 2 hours at 4°C. Nuclei with counterstained with DAPI before mounting the slides. No staining was performed after cryosectioning Tg(isl2b:GFP)zc7. The following primary antibodies were used: Rabbit cleaved Caspase3 (Asp175) (1:250, Cell Signalling Technologies), mouse HuC (1:100,

Life Technologies-Novex), mouse anti-PCNA (1:100, ZYMED laboratories), rabbit anti-

Phospho-histone H3 (Ser10) (1:250, Cell Signalling Technologies), rabbit anti-Pax6

(1:100, Covance), rabbit anti-PKC (1:100, Santa Cruz Biotechology, Inc.), mouse anti-

Zn5 (1:100, ZIRC), and mouse anti-Zpr1 (1:200, ZIRC). Secondary antibodies used for detection were from Jackson ImmunoResearch Laboratories: mouse, rat and rabbit Cy3

(1:500), mouse and rabbit Cy5 (1:200). Images were taken using a Leica TCS Sp5 II

Confocal Microscope and analyzed with Leica LAS AF software. For cleaved caspase 3 positive cell counts, total numbers averaged from three sections derived from different embryos of each group were used. Statistical analyses between MO-injected and control- injected groups were performed using Student’s t-test.

3.3.10 Chromatin immunoprecipitation (ChIP)

ChIP experiments were performed in duplicate using cells derived from ~100 zebrafish heads at 48 hpf. Head cells were cross-linked using 1% formaldehyde for 8 min and quenched using 0.125M glycine. Cells were then centrifuged at 470 x g for 10 min at

4°C and resuspended in 500ul ice-cold PBS supplemented with protease inhibitors

(Roche). Sonication was performed using Bioruptor (Diagenode) using 7 cycles of 30s

“ON”, 30s “OFF”. ChIP was performed using the Millipore EZ-ChIP kit according to the manufacturer protocol. ChIP-grade antibodies for anti-H3K9me2 (AbCam) and an

isotype control (AbCam) were used at 2µg per sample. The promoter regions from vsx2 and ccnd1 were amplified by PCR (35 cycles) using the primers listed in Table 3.4.

Table 3.4. Primers sequences used to PCR amplify the promoter regions of zebrafish ccnd1 and vsx2.

Gene Forward primer (5’ à 3’) Reverse primer (5’ à 3’) ccnd1 (-424 to -15) TTCACCCCAGTCTTTTCCAC AAAGTCTCGCTGCAGCTCTC

vsx2 (-478 to -67) AGACATTTTCCAGCGCACTT GCAATACGCATGATCCCTCT

3.4 Results

3.4.1 The expression of znf644 paralogues during embryogenesis

Most HMT-related proteins are widely conserved across metazoa (Pu et al.,

2010), allowing experimental evaluation of function using appropriate model organisms.

I exploited this principle to investigate the developmental function(s) of ZNF644 vis-à- vis G9a during zebrafish embryogenesis, an experimentally tractable system particularly with regards to the developing nervous system and retina. It has been reported that zebrafish g9a is broadly expressed during embryogenesis and that disruption of its function via morpholino injection causes severe morphological defects, notably reductions in eye and brain size (Rai et al., 2010). Using human ZNF644 for a BLASTP search against the reference zebrafish genome (Zv9), I identified two putative ZNF644 orthologues, which I termed znf644a and znf644b and which showed 27.1% and 19.6%

amino acid identity to human ZNF644, respectively. Like ZNF644, znf644a and znf644b contain a highly conserved atypical C2H2-like ZF motif in their C-terminal regions, implying that the physical association with G9a/GLP is likely also conserved (Refer back to Figure 2.8). Conversely, the paralogues showed differential retention of a reduced complement of C2H2-like ZF motifs; in other words, znf644a lacks ZF3 and ZF4, while znf644b lacks ZF1 and ZF5) (Figure 3.1), suggesting the possibility of sub- functionalization between the paralogues.

1 2 3 4 5 6 7

Figure 3.1: Protein architecture of human ZNF644 and zebrafish paralogues Znf644a and Znf644b. The relative positioning of the C2H2-like and atypical C2H2-like ZF motifs within these proteins in are indicated. Note the absence of ZF3 and ZF4 in

Znf644a and ZF1 and ZF5 in Znf644b.

To identify the physiological context(s) in which the G9a-ZNF644 physical association may exert its function, I first profiled the embryonic expression pattern of znf644a and znf644b using in situ hybridization (ISH; Figure 3.2). In stark contrast to the broad expression pattern of g9a (Rai et al., 2010), the expression of both znf644 paralogues was highly spatially restricted to RPCs, the emerging retinal ganglion cell

(RGC) layer, and in discrete domains of the midbrain. More specifically, znf644a was prominently found in the emergent dorsal midbrain (presumptive tectum) and further into the tectal lobes, while znf644b was predominantly localized to ventricular zone progenitor cells in the developing dorsal and ventral midbrain (presumptive tectum and tegmentum) (Figure 3.3). Not only was the expression of the two paralogues spatially confined to these regions, but also constrained temporally at a time point at which retinal and neural progenitor cells are on the verge of undergoing cell cycle withdrawal and differentiation (Agathocleous and Harris, 2009). The overlapping expression of znf644a and znf644b in progenitor cells of the retina and midbrain suggested the possibility of functional cooperatively with g9a in the developing retina and midbrain.

Figure 3.2: Expression pattern of znf644a and znf644b during zebrafish embryogenesis. Whole-mount in situ hybridization experiments monitoring the expression (blue stain) of (A) znf644a (n=15) and (B) znf644b (n=15) mRNA at the indicated developmental stages or time points. Red lines highlight midbrain regions.

Figure 3.3: Expression of znf644a and znf644b in neural progenitor populations of the developing zebrafish retina and midbrain at 24 hpf. Representative lateral, retina cross- section and midbrain cross-sections of whole-mount in situ hybridization experiments assaying the expression of znf644a or znf644b at 24 hpf (n=15 in each group). Red arrows highlight expression in progenitors in the presumptive ganglion cell layer of the retina. Red brackets indicate expression in midbrain regions.

3.4.2 Disruption of G9a or ZNF644 causes defects in the forming retina and midbrain

To examine the functional importance of znf644a and znf644b in relation to g9a, I applied a genetic loss-of-function approach using a previously validated splice-block morpholino (MO) targeted against g9a (g9a-MO) and controlling for sequence specificity using a mismatch control (Rai et al., 2010). It is important to emphasize that the use of splice-block MOs do not generally affect gene function in the early embryo (i.e., blastulation, gastrulation, etc.) because they do not target pre-spliced maternally-supplied mRNA. Hence, the phenotypes caused by MO injection generally reflect gene functions in later developmental stages and in organogenesis. As neither znf644a nor znf644b had been previously studied in zebrafish, I designed splice-block MOs targeted against znf644a (znf644a-MO) and znf644b (znf644b-MO) as well as corresponding mismatch controls. Consistent with previous reports (Rai et al., 2010), injection of g9a-MO in early

(1-2 cell) embryos induced broad and severe morphological abnormalities, including reductions in eye and brain size, in a dose-dependent manner (Figure 3.4). This is consistent with the idea that g9a is a critical developmental regulator in multiple tissue contexts (see Table 1.4), although the precise cellular and molecular defects underlying the g9a morphant phenotype(s), such as the apparent defects in eye and midbrain development, were unknown.

Relative to the severe morphological defects observed in g9a morphant embryos, the morphological abnormalities of znf644a and znf644b morphant embryos were generally mild – specifically, these morphants exhibited significant reductions in eye size and altered midbrain morphology (i.e., reduced midbrain size in znf644a morphants and

expanded midbrain in znf644b morphants) (Figure 3.5). Mismatch control MOs showed no obvious defects (data not shown), demonstrating that these morphological defects are unlikely to stem for non-sequence-specific targeting defects. Moreover, complementation experiments based on co-injection of the in vitro transcribed, capped cognate mRNA rescued the morphological defects of znf644a and znf644b morphants (Figure 3.5), further validating targeting specificity.

As ZNF644 mutations were linked to high-grade myopia (Shi et al., 2011; Tran-

Viet et al., 2012), I focused on the microphthalmia (small eye) phenotypes of g9a, znf644a and znf644b morphants. To this end, I optimized the MO injection doses for each targeted gene so as to result in an approximate 50% reduction in the 2D retinal cross-sectional area (Figure 3.5), corresponding to a near complete splicing disruption of znf644a, an approximate half (57%) decrease in mature znf644b, and an approximate one-third (35%) decrease in g9a levels (Figure 3.6 and data not shown). These data highlighted the forming retina and midbrain as neural structures that are critically dependent on g9a, znf644a and znf644b function for their proper development.

Normal Mildly abnormal Abnormal Severely defected

Normal

Mildly abnormal

Abnormal

Severely abnormal

Figure 3.4. The varying morphological defects of g9a morphant embryos. (A-D) The varying degree of development defects observed in g9a morphant retinas. n=5 in each case. (E) Quantitative measurement of the frequency at which the developmental defects occurred as a function of g9a-MO or g9a-mMO (mismatch control) injection dose.

A B C D

Figure 3.5: Reduced eye size and altered midbrain morphology in g9a, znf644a and znf644b morphant embryos. (A-D) Lateral views of WT (21/21), g9a (20/32), znf644a

(14/21), and znf644b (10/18) morphant embryos. Red dotted circles indicate the approximate retina 2-dimensional area of the WT retina. Red arrowheads highlight midbrain regions. (E) Quantitative measurements of retinal cross-sectional areas, in

µm2, of g9a, znf644a, and znf644b morphant embryos. Asterisks indicate p-values < 10E-

19. (F-J) Lateral views of WT (36/36), znf644a morphants (23/35), znf644a morphants with znf644a mRNA co-injected (10/19), znf644b morphants (14/25), and znf644b morphants with znf644b mRNA co-injected (7/8). (K) Summary of the frequency at which the morphant “defective” or WT (“normal”) phenotypes were observed in the indicated embryos.

unspl znf644a spl

znf644b unspl spl β-actin

WT MO MO - - znf644a znf644b

Figure 3.6. MOs targeted against znf644a or znf644b result in transcript-specific splicing defects. RT-PCR experiments monitoring the steady state levels of unspliced

(unspl) and spliced (spl) transcripts of znf644a or znf644b in the indicated MO treated embryos at 24 hpf. RNA was prepared from dechorionated whole embryos (see Methods).

3.4.3 Severe cell viability and proliferative defects are observed in g9a morphant retinas

While previous characterization of the zebrafish g9a morphant phenotype had reported morphological defects in eye and midbrain development similar to those I observed (see Figures 3.4-3.5) (Rai et al., 2010), the cellular defects underlying the small eye phenotype of g9a morphant retinas were not fully investigated. To better understand the g9a morphant phenotype, I performed various immunostaining and in situ hybridization assays using markers of apoptosis, proliferation, and differentiation, as aberrant regulation of these processes could account for the observed hypocellularity and reductions in eye size. The following sections will describe these assays and reports the key findings.

First, I performed immunostaining assays using retinal cross-sections from g9a morphants to monitor levels of Caspase3 activation. While there are p53-independent cell death pathways (Tedeschi and Giovanni, 2009) as well as there is cell-death independent caspase function (Fernando et al., 2005), the small eye phenotypes helped to point us in the direction that either cells are not proliferating or cells are dying. Based on the published literature (Prykhozhij, 2010), increased caspase3 activation is strongly correlated with cell death in the zebrafish retina. Indeed, I found strikingly high levels of apoptotic cells throughout the retinas of g9a morphants at both 48 and 72 hpf (Figure

3.7A-B), suggesting a critical role for G9a in maintaining the viability of retinal cells.

This is consistent with previous indications that G9a function is closely related to cell survival in various contexts (i.e., early mouse embryos (Tachibana et al., 2002) and ethanol-induced embryonic neurodegeneration (Subbanna et al., 2013). However, the

cellular mechanisms and pathways related to G9a-mediated regulation of survival are unclear. Apoptosis can occur through either extrinsic pathway activation (when cells receive external signals that activate death receptors) or intrinsic (mitochondrial) pathway activation (when pro-apoptotic factors are released from mitochondria after their permeablization), and activation of the p53 pathways is the primary driver of the intrinsic apoptotic pathway (Michalak et al., 2005). To determine whether p53 activation mediated the elevated levels of cell death in the g9a morphant retina (and hence implicating the intrinsic pathway), I used a MO previously shown to effectively block p53 protein translation (p53-MO) during zebrafish development (Nasevicius and Ekker, 2000). I found that co-injection of p53-MO led to a partial, yet statistically significant, rescue of the apoptotic phenotype (Figure 3.7A-B). This suggests that the broad apoptotic phenotype (i.e., Caspase3 activation) is at least partially driven by the intrinsic p53- medated cell death pathway.

Figure 3.7: Impaired cell viability in g9a morphant retinas. (A) Immunostaining experiments monitoring the levels of activated Caspase3 in retinal cross-sections at 48 and 72 hpf from WT, g9a morphants, and g9a morphants in which p53-MO was co- injected (n=3 for each group). Yellow arrows indicate activated Caspase3-positive cells.

(B) Quantitative measurements of the number of activated Caspase3-positive cells in retinal cross-sections at 72 hpf in WT, g9a morphants, and g9a morphants with p53-MO co-injected. Error bars represents standard deviation. *p-value < 0.05; **p-value <

0.005; ***p-value < 0.005. Student’s T-test.

In addition to apoptosis, I assessed the proliferative state of g9a morphant retinal cells at 48 hpf using immunostaining assays monitoring the incorporation of BrdU as well as the levels and localization of phospho-histone H3 (pHH3)-positive and PCNA- positive cells. At 48 hpf, proliferative cells are normally confined to a progenitor cell niche known as the ciliary marginal zone (CMZ) and are mostly absent from the central

retina (see Figure 3.8A-B). I observed that g9a morphant retinas showed increased levels of BrdU incorporation, pHH3-positive cells as well as PCNA-positive cells in the central retina. Hence, in addition to elevated levels of apoptosis, g9a morphant retinas also exhibit persistent cellular proliferation in the central retina at 48 hpf.

A B

Figure 3.8. G9a morphant retinas exhibit increased cellular proliferation. (A)

Immunostaining assays using retinal cross-sections from WT or g9a morphants monitoring the levels of BrdU incorporation, pHH3 and PCNA at 48 hpf. The yellow box highlights the central retina as well as the area used for quantitation (n=3). (B)

Quantitative measurements of BrdU-positive and pHH3-positive cells. Error bars reflect standard deviation. * p < 0.05, Student’s T-test.

Given the elevated levels of apoptosis and evidence of persistent cellular proliferation in g9a morphant retinas, I performed immunostaining assays to determine

how these cellular defects affected progenitor and/or differentiated cells populations at 48 and 72 hpf. I found that the retinas of g9a morphant embryos largely lacked differentiated neurons (e.g., GABA-positive amacrine cells, PKC-positive bipolar cells, and Zpr1- positive cone photoreceptors) at 72 hpf (Figure 3.9). In addition, g9a morphant retinas lacked Vsx2-positive cells at 48 hpf in the CMZ, which is characteristic of fully multipotent RPCs that normally reside in that region (Figure 3.9). In addition, g9a morphants also lacked Vsx2-positive cells were in the central retina, which marks a subset of bipolar neurons (Figure 3.9). This data suggests that g9a morphant retinas not only lack various types of differentiated cells, but may also be depleted of fully multipotent progenitor cells at 48 and 72 hpf.

Figure 3.9. G9a morphant retinas lack progenitor cells as well as various types of differentiated neurons. Representative immunostaining assays using retinal cross

sections from WT or g9a-MO embryos monitoring the expression of the indicated protein marker at the indicated time point (n=3 in each group). Vsx2-positive cells mark progenitors (green arrows) as well as a subpopulation of bipolar cells. GABA marks horizontal cells, PKC marks bipolar cells, and Zpr1 marks photoreceptor cells. Yellow arrows point to populations of differentiated cells.

Midway through my characterization of the g9a morphant retina, the role of G9a in mouse RPCs via conditional knockout was published (Katoh et al., 2012). Much like I had observed in zebrafish g9a morphants, G9a knockout in RPCs resulted in strikingly elevated levels of apoptosis. However, in contrast to the g9a morphant retina phenotype I had observed, they also detected the formation of various types of differentiated neurons, albeit in reduced numbers relative WT retinas. Importantly, they used RNA microarray and ChIP assays to identify a set of RPC-specific genes that were silenced by G9a- mediated H3K9me2 in their promoter regions. These genes include certain master retinal

TFs, such as Vsx, Hes1, Lhx2, as well as various types of cyclins. Among these RPC- specific genes targeted by H3K9me2, Vsx2 and CyclinD1 have particularly well characterized functional homologues in zebrafish (vsx2 and ccnd1, respectively) that are reported to perform similar regulatory roles as their mammalian counterparts in the developing retina. I therefore performed in situ hybridization experiments to determine whether these genes were upregulated in g9a morphant retinas at 48 hpf (a time point at which their expression is normally shutoff in the central retina). However, I observed only modest changes (if any) in the expression of vsx2 and ccnd1 relative to control embryos at 48 hpf (Figure 3.10). It was not initially obvious whether the lack of obvious

changes in vsx2 or ccnd1 expression levels in g9a morphants indicated that (1) g9a was not required for the suppression of vsx2 and ccnd1 in the zebrafish retina, that (2) sufficient g9a activity remained in g9a morphant retinas to suppress the expression of vsx2 and ccnd1, or that (3) cells that would otherwise normally exhibit mislocalized expression of vsx2 and ccnd1 had already already depleted (see Figure 3.9). At this point in the project, it was therefore unclear whether or not zebrafish g9a actually played a role in suppressing vsx2 and ccnd1, as had been reported in mouse RPCs. It was not until characterization of znf644a and znf644b that additional insights regardin the role of

G9a/H3K9me2 in suppressing vsx2 and ccnd1 in the zebrafish retina became apparent.

Figure 3.10. Expression of vsx2 and ccnd1 in the g9a morphant retina at 48 hpf. In situ hybridization assays monitoring the expression of vsx2 or ccnd1 in retinal cross-sections derived from WT or g9a morphant embryos at 48 hpf. Red arrows highlight the CMZ regions of the retinas. n=5 in each case.

3.4.4 Znf644a and znf644b morphant retinas both exhibit mislocalized expression of vsx2 and ccnd1 and reduced promoter H3K9me2

Following my characterization of the g9a morphant retina, I sought to use these same cellular and biochemical assays to investigate the functions of the zebrafish znf644 paralogues. My initial hypothesis was that disrupting the function of znf644a or znf644b might give rise to retinal phenotypes similar to those I described in g9a morphant retinas.

This, however, proved not to be the exact case, as disrupting the function of either paralogue gave rise to phenotypes that not only differed from each other, but also from g9a morphants, as will be described here.

First, I used in situ hybridization experiments to monitor the expression of vsx2 and ccnd1 at 48 hpf to see whether these transcripts would show persistently upregulated expression. Indeed, I found that both znf644a and znf644b morphant retinas showed strong mislocalized expression of vsx2 and ccnd1 transcripts throughout the central retina at 48 hpf, suggesting that both znf644a and znf644b are required to suppress the expression of these RPC-specific genes (Figure 3.11A). In addition to vsx2 and ccnd1, I assayed the expression of various other progenitor and proneural genes to assess whether znf644a and znf644b morphant retinas represented To determine whether the observed mislocalized expression of vsx2 and ccnd1 was due to defects in promoter H3K9me2, I

performed chromatin immunoprecipitation (ChIP) assays from chromatin preparations derived from embryonic head region cells at 48 hpf. Indeed, I found that H3K9me2 normally marked the promoter regions of vsx2 and ccnd1 at 48 hpf and that this methylation mark was significantly reduced in znf644a and znf644b morphant retinas

(Figure 3.11B). These data suggest that znf644a and znf644b are in vivo regulators of the

H3K9me2 mark at the promoters of key RPC-specific genes.

To determine whether the mislocalized expression of vsx2 and ccnd1 in znf644a and znf644b morphant retinas simply represented a failure to silence vsx2 and ccnd1 expression or whether it signified a more drastic switch in cell identity and behavior, I examined the expression of additional multipotency (e.g., rx3) and proneural (e.g., ath5)

TFs that are used to distinguish RPCs from other types of retinal cell types. In addition to the mislocalized expression of vsx2 and ccnd1 throughout the central retina, I found that retinal cells of znf644b morphants also exhibited elevated and persistent expression of a host of multipotency-related and proneural-related transcripts (Figure 3.11A and data not shown), suggesting that these cells express several genes that typify an RPC-specific gene expression. In contrast, other than vsx2 and ccnd1, I observed no significant changes in expression for multipotency or proneural genes in znf644a morphant retinas at

48 hpf, suggesting the dysregulated expression of a subset of RPC-specific genes.

Figure 3.11. Znf644a and znf644b morphant retinas exhibited mislocalized expression of vsx2 and ccnd1 as well as reduced H3K9me2 levels in the vsx2 and ccnd1 promoter regions. (A) In situ hybridization assays monitoring the expression of the progenitor

genes vsx2, rx3 and ccnd1 as well as the proneural factor ath5 at 48 hpf in WT, znf644a, and znf644b morphant retinas (n=3). (B) ChIP-PCR assays using chromatin prepared from whole head embryonic head region cells (~100 embryos) at 48 hpf. An anti-

H3K9me2 antibody or an isotype control (IgG) antibody was used for ChIP, and the indicated promoter regions were amplified by PCR.

3.4.5 Znf644b morphant retinas maintain a fully multipotent proliferative state, whereas znf644a morphant retinas generate differentiated cells

In light of the mislocalized expression of vsx2 and ccnd1, I hypothesized znf644a and znf644b morphant retinas may be able to maintain a fully multipotent proliferative state for a prolonged period of time. To this end, I assessed the proliferative state of znf644a and znf644b morphant retinas at 48 hpf using a BrdU incorporation assay as well as immunostaining for pHH3 and PCNA levels. Indeed, both znf644a and znf644b morphant retinas exhibited increased numbers of BrdU-positive, pHH3-positive, and

PCNA-positive cells in the central retina (Figure 3.12A-B), suggesting prolonged proliferative capacity cells of the central retina. However, I also observed that there were considerably more pHH3-positive cells in znf644b morphant retinas relative g9a or znf644b morphant retinas, which may indicate more rapid progression through the cell cycle.

A B

Figure 3.12. Persistent proliferation in znf644a and znf644b morphant retinas. (A)

Immunostaining assays using retinal cross-sections monitoring BrdU incorporation, pHH3-positive cells, and PCNA-positive cells at 48 hpf in WT, znf644a morphants or znf644b morphant (n=3 for each group). (B) Quantitative measurement of the number of

BrdU-positive or pHH3-positive cells in the central retina. The yellow box highlights the central retina and the approximate counting area for quantitation.

As I had done with g9a morphant retinas, I performed immunostaining assays to determine whether the persistent proliferation in znf644a and znf644b morphant retinas affected the formation of differentiated types of neurons. Given the persistent expression

of vsx2 and ccnd1 throughout the retina at 48 hpf, one might anticipate a maintained proliferative state and a failure to differentiate. However, as a population, cells of znf644a morphant retinas were nonetheless able to form various types of differentiated neurons at 48 hpf, including bipolar neurons, horizontal cells and photoreceptors (Figure

13). Thus, the znf644a morphant retinal cells undergo differentiation despite increased levels of cellular proliferation and the mislocalized expression of vsx2 and ccnd1. In striking contrast to cells of znf644a morphant retinas (described above), znf644b morphant retinas were largely void of GABA-positive horizontal cells, PKC-positive bipolar cells and Zpr1-positive cone photoreceptors (Figure 3.13). Instead, znf644b morphant retinas were composed of Vsx2-positive cells throughout the retina at a time when this type of cell is normally confined to the ciliary marginal zone (CMZ) (Figure

3.13). These data suggest that znf644b morphant retinas maintain a prolonged proliferative progenitor identity.

Figure 3.13. Znf644a and znf644b morphant retinas are composed of different types of cells. Immunostaining assays from retinal cross-sections from WT, znf644a morphant or znf644b morphant embryos monitoring the expression of the indicated protein marker at the indicated time point (n=3 for each group). Vsx2-positive cells at 48 hpf mark progenitor cells of the CMZ (grey arrow) or a subpopulation of bipolar neurons (yellow arrows). GABA marks horizontal cells, PKC marks bipolar cells, and Zpr1 marks cone photoreceptors.

Incidentally, it has been well demonstrated that misexpression of progenitor- specific genes, such as vsx2 and ccnd1, in differentiating or differentiated neuronal cells

is typically sufficient to induce an apoptosis phenotype (Bonda et al., 2009; Zhu et al.,

2004); unpublished data from the Tropepe lab). Hence, I hypothesized that differentiated neuronal populations of znf644a morphant retinas may likewise exhibit impaired cell viability. Indeed, I found that the differentiated regions of znf644a morphant retinas exhibited a highly temporally and spatially confined activation of Caspase3 (Figure

3.14). It is important to note that this apoptosis phenotype occurred in mature populations of differentiated cells, in contrast to the g9a morphant retina where apoptosis occurred earlier in post-mitotic progenitors and/or early differentiating cells (Figure 3.7). Also, in contrast to the apoptotic phenotype of g9a morphant retinas that was partially rescued by co-injection of p53-MO, Caspase3 activation in the znf644a morphant retina was much more efficiently rescued by co-injection of p53-MO (Figure 3.14), indicating that apoptosis in the znf644a morphant retina is primarily driven via the p53 pathway. In contrast to znf644a morphants, znf644b morphant retinas showed no detectable activation of Caspase3 at 48 or 72 hpf (Figure 3.14).

Figure 3.14. Znf644a morphant retinas, but not znf644b morphant retinas show elevated levels of apoptosis in mature populations of differentiated neurons. (A)

Immunostaining assays using retinal cross-sections from WT, znf644a morphants or znf644b morphants monitoring the activation of Caspase3 at 48 or 72 hpf. The yellow arrow highlights Caspase3+ cells (n=3 for each group). (B) Quantitative measurements of the number of Caspase3-positive retinal cells in WT, znf644 morphants, or znf644 morphants with p53-MO co-injected. Error bars reflect standard deviation, and the asterisks indicates p < 0.05, Student’s T-test.

Because the znf644b morphant retinas maintained the expression of RPC-specific genes all the while maintaining a proliferative state throughout the, I hypothesized that these retinal cells may exhibit delayed differentiation. To this end, I performed additional

immunostaining assays to monitor the formation of the first-born retinal subtypes, isl2b- positive retinal ganglion cells (RGCs) and pax6a-positive amacrine cells. Indeed, whereas

RGCs and amacrine cells have formed in WT retinas by 48 hpf, these types of cells were lacking in znf644b morphant retinas at this time point, but rather began to emerge at the

72 hpf time point (Figure 3.15). It is unlikely that the neuronal birth order was altered

(i.e., RGCs and amacrine cells “skipped) because HuC, which marks various types of differentiated retinal cells, was lacking outside of the emerging RGCs and amacrine cells at 72 hpf (Figure 3.15). Collectively, this data suggests that znf644 morphant retinas exhibit developmental delays that include a prolonged proliferative progenitor identity and the delayed entry into differentiation.

Figure 3.15: Delayed differentiation in znf644b morphant embryos. Immunostaining experiments monitoring the expression of Pax6, Isl2b, or HuC in retinal cross sections at

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the indicated time points (n=3). Pax6 marks amacrine cells and a subset of RGCs, Isl2b marks a subset of RGCs, and HuC marks various types of post-mitotic neuronal cells.

3.4.6 Summary of g9a, znf644a and znf644b morphant phenotypes in the retina

The retinal phenotypes of g9a, znf644a, and znf644b did exhibit some similarities; however, unique distinguishing features were evident from the assays I performed to assess the cellular proliferative capacity, gene expression patterns, differentiation state and apoptosis of these retinal cells. A summary of the individual morphant phenotypes is provided in Table 3.5 to enable a more straightforward comparison between the morphant retinal cell behaviors and characteristics.

Table 3.5. Summary of the g9a, znf644a and znf644b morphant retinal phenotypes based on the results from in situ hybridization and immunostaining assays described earlier in this Chapter. The Comments column summarizes the physical and behavioral characteristics of the retinal cells of the indicated morphants.

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3.4.7 H3K9me2-positive retinal nuclei are visible in differentiated cells.

In addition to markers of multipotency, cell cycle, differentiation and apoptosis, I also investigated global levels of H3K9me2 in retinal cells using immunostaining. Cells with H3K9me2-positive nuclei had been observed in cross-sections the mouse embryonic retina (Rao et al., 2010), although it is currently unknown when these H3K9me2-marked nuclei are formed within the context of retinal differentiation. I found that H3K9me2- positive nuclei were uniquely visible later in the later stages of retinal differentiation among differentiated cells of the ganglion and inner nuclear layers at 72 hpf (Figure

3.16). Note that znf644a and znf644b are specifically and transiently expressed at RPCs at 24 hpf (See Figure 3.3), and it is thus highly unlikely that these H3K9me2-positive nuclei are directly regulated by znf644a and znf644b. It is possible that this late temporal emergence of H3K9me2 represents the aforementioned expanded regions of H3K9me2- marked chromatin (Chen et al., 2012) that are perhaps initially “seeded” by the earlier activity of znf644a and/or znf644b in differentiating RPCs.

I then sought to determine whether disruption of g9a, znf644a, or znf644b affected the formation of H3K9me2-positive nuclei at 72 hpf, I used immunostaining to examine the formation of H3K9me2-postive nuclei. I found a reduced amount, but not a complete loss, of H3K9me2-postive nuclei in g9a morphant retinas (Figure 3.16).

Because G9a has been reported to be the primary H3K9me2 HMT in the retina (Katoh et al., 2012), the partially reduced H3K9me2 staining in these retinas may reflect partial reductions in g9a activity in g9a morphant retinas. Similar to g9a morphant retinas, I also observed slightly fewer H3K9me2-postiive nuclei in differentiated retinal cells in the

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znf644a morphants relative WT (Figure 3.16), suggesting that znf644a activity at 24 hpf has little effect on the formation of H3K9me2-positive nuclei at 72 hpf (Figure 3.16).

Interestingly, the znf644b morphant retina uniquely showed an absence of H3K9me2- positive nuclei at 72 hpf (Figure 3.16). Recall that znf644b morphant retinas maintain a prolonged proliferative progenitor identity and undergo differentiation later than normal retinal cells (see Figure 3.15), suggesting that znf644b indirectly controls the formation of H3K9me2-positive nuclei at 72 hpf by preventing delaying differentiation and enabling for properly expanded H3K9me2-marked chromatin at the 72 hpf.

Figure 3.16. H3K9me2-positive nuclei are visible in differentiated cell populations.

Immunostaining experiments using retinal cross-sections from WT, g9a morphants, znf644a morphants, and znf644b morphants monitoring H3K9me2 levels at 72 hpf (n=3).

Yellow arrows point to H3K9me2-positive cell populations.

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3.4.8 Extensive functional cooperativity between g9a and the znf644 paralogues

Due to seemingly non-overlapping aspects of the phenotypes of g9a, znf644a and znf644 morphant retinas, I sought to determine the functional relationship between g9a, znf644a and znf644b using a “genetic cooperativity” assay. In this approach, subphenotypic injection doses of MOs, which alone introduce partial splicing deficiencies that are insufficient to yield significant developmental defects (Figure 3.17 and data not shown), were co-injected to reveal potentially additive or synergistic effects.

Conceptually, if two proteins are components of the same complex, the double MO treatment would constitute a “double-hit” on this complex, perhaps resulting in sufficiently impaired complex function so as to disrupt the regulatory function of the complex and give rise to a detectable phenotype. Such an approach has previously been used in zebrafish to demonstrate the functional dependency/cooperativity between G9a and the DNA methylation enzyme DNMT3 (Rai et al., 2010).

3.4.8.1 G9a and znf644a cooperatively regulate the viability of differentiating retinal neurons

Co-injected subphenotypic doses of g9a-MO and znf644a-MO gave rise to an apoptotic response similar to that found in higher-dose g9a morphant retinas – that is, a relatively early and broad cell death phenotype in various cell types throughout the retina

(Figure 3.17A). There were, however, some persistently elevated levels of pH3 (Figure

3.17C), perhaps indicating a failure of mitotic cells to fully withdraw from the cell cycle.

The co-injection of subthreshold MO doses did not appreciably affect the expression pattern of vsx2 (Figure 3.17D), again consistent with the phenotype of the full-dose g9a

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morphant retina. These data suggest that g9a and znf644a function cooperatively to suppress apoptosis in progenitors and differentiating retinal cells.

Figure 3.17: Extensive functional cooperativity between g9a, znf644a and znf644b. (A)

Immunostaining experiments monitoring the levels of activated Caspase3 in the retinas of embryos injected with individually subphenotypic (“low”) doses of g9a-MO, znf644a-MO or znf644b-MO as well as combination subphenotypic doses of g9a-MO together with znf644a or znf644b (n=3 in each group). (B-C) Immunostaining experiments monitoring the levels of PCNA or pHH3 in the retinas of embryos injected with subphenotypic doses

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of g9a-MO, znf644a-MO or znf644b-MO as well as co-injection of the indicated subphenotypic (“low”) doses (n=6 in each group). (D) Whole-mount in situ hybridization experiments monitoring the expression of vsx2 in the retinas of embryos injected with subphenotypic (“low”) doses of g9a-MO (n=11), znf644a-MO (n=10) or znf644a (n=0) as well as co-injection of g9a-MO and znf644a-MO (n=13), g9a-MO and znf644b-MO

(n=10) and znf644a-MO and znf644b-MO (n=10).

3.4.8.2 G9a and znf644b cooperatively suppress a prolonged proliferative state in the retina

More notably, co-injection of subphenotypic doses of g9a-MO and znf644b-MO, led to markedly increased levels of cell proliferation markers (Figure 3.17B-C), comparable to those observed in the full dose znf644b morphant retinas, yet again without affecting cell viability (Figure 3.17A). Intriguingly, despite the prolonged proliferation phase evident throughout most of the retina, there were again minimal changes to the expression pattern of vsx2 in these morphants (Figure 3.17D) This was surprising, given that multipotency-related TFs (such as vsx2) are well-established positive upstream regulators of proliferation in expanding populations of RPCs (Green,

Stubbs and Levine, 2003; Vitorino et al., 2009). However, the extended maintenance of a proliferative state, despite the suppressed expression of multipotency-related TFs (such as vsx2), implies a mechanistic dissociation between cell cycle progression and the presence of multipotency-related master regulators in later populations of RPCs. My data also suggest that, normally, g9a functions cooperativity with znf644b to suppress a prolonged proliferative state.

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3.4.8.3 Znf644a and znf644b cooperatively suppress a prolonged proliferative RPC state in the retina

In addition to investigating “genetic cooperativity” between g9a and each znf644 paralogue, I investigated the possibility that the two znf644 paralogues exhibit some degree of functional cooperativity. The retinas of znf644a and znf644b morphants both showed mislocalized expression of vsx2 and ccnd1, as described in section 3.4.4, and the functions of the two genes appear to be overlapping in this regard. Indeed, co-injection of subphenotypic doses of znf644a-MO and znf644b-MO likewise resulted in mislocalized expression of vsx2 throughout most of the retina (Figure 3.17D) along with an increased number of proliferative and mitotic cells (Figure 3.17B-C). These data are consistent with an overlapping and cooperative function for znf644a and znf644b in transitioning proliferating RPCs towards differentiated post-mitotic progeny.

3.4.9 The functions of human ZNF644 encompasses those of both zebrafish paralogues

In contrast to zebrafish, which, according to the data I have presented here, express two paralogous znf644 genes that function cooperatively with each other and with G9a, mammalian genomes encode a single version of ZNF644. I wondered if its function more closely reflects the function of one particular paralogue, or rather possessed the functionality encapsulated by both paralogues? I therefore assessed the functional conservation between human ZNF644 and the zebrafish orthologues by performing functional complementation experiments using mRNA encoding the human version of ZNF644. Strikingly, I found that native full-length ZNF644 equally efficiently rescued the retinal defects of both znf644a and znf644b morphant retinas; namely, reversing the mislocalized expression of vsx2 (Figure 3.18A) and the elevated levels of

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apoptosis in znf644a morphants (Figure 3.18B), and the mislocalized expression of vsx2

(Figure 3.13C) and the prolonged proliferative phase (Figure 3.13D) and delayed differentiation (data not shown) in znf644b morphant retinas. Hence, the function of human ZNF644 likely encompasses those of both of the zebrafish paralogues, suggesting conserved functionality in mammalian systems.

Figure 3.18. Human ZNF644 rescues the znf644a and znf644b morphant phenotypes, but requires physical association with G9a/GLP. (A) Whole-mount in situ hybridization assays monitoring the expression of vsx2 at 48 hpf in znf644a morphant retinas showing rescue by co-injection of mRNA encoding WT human ZNF644 (n=14, 50% normal) but not the C1263A mutant version (n=9, 44% normal). (B) Immunostaining experiments monitoring the activation of Caspase3 at 72 hpf in znf644a morphant retinas after co- injection of mRNA encoding WT human ZNF644 (n=3) or the C1263A mutant (n=3). (C)

Whole-mount in situ hybridization assays monitoring the expression of vsx2 at 48 hpf in znf644a morphant retinas rescued by co-injection of WT human ZNF644 mRNA (n=12,

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83% normal) compared to the C1263A mutant version (n=9, 22% normal). (D)

Immunostaining assays monitoring the expression of PCNA at 48 hpf in znf644b morphant retinas after co-injection of WT human ZNF644 (n=3) or the C1263A mutant

(n=3).

I then sought to determine whether the physical association with G9a and GLP I documented in human was likewise important for the function of ZNF644 in zebrafish.

To this end, I evaluated the C1263A and H1283A mutant alleles I had generated, which, as I showed earlier, failed to co-purify G9a and GLP (Figure 2.8). Indeed, these same mutants failed to rescue any of the described retinas defects of full dose znf644a or znf644b morphants (Figure 13.18A’-D’ and data not shown). These data suggest the function of ZNF644 is both evolutionarily conserved and critically dependent on its ability to bind to G9a (and GLP).

3.4.10 The retinal defects in G9a and ZNF644 morphants are recapitulated in the midbrain

Due to the temporally parallel retina- and midbrain-specific expression of znf644a and znf644b, I examined whether the cellular defects of g9a, znf644a and znf644b morphant retinas were recapitulated in the developing midbrain. First, I found that the midbrains of znf644a and znf644b morphants exhibited strong mislocalized expression of ccnd1 at 48 hpf, as was also observed in the midbrain (Figure 3.19A). Also the midbrain regions in which the mislocalized expression of ccnd1 occurred were in similar regions that normally express znf644a or znf644b earlier at 24 hpf (compare with Figure 3.3). In addition, similar to the retinal defects of g9a morphants, g9a morphant midbrains showed

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elevated levels of Caspase3 activation throughout the midbrain at 72 hpf, which was similarly partially rescued by co-injection of p53-MO (Figure 3.19B). Increased apoptotic cells were also evident in znf644a morphant midbrains at 72 hpf, and, as in the retina, this apoptosis was efficiently rescued by co-injection of p53-MO (Figure 3.19B).

In addition, znf644b morphant midbrains similarly exhibited increased PCNA+ proliferative cells throughout the midbrain (Figure 3.19C), as was also seen in the retina.

Collectively, these data suggest that the functions of g9a, znf644a and znf644b in RPCs are likely recapitulated in midbrain progenitor cells.

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Figure 3.19. The defects in gene expression, proliferation and survival in g9a, znf644a and znf644b morphant retinas are recapitulated in the midbrain. (A) In situ hybridization assays monitoring the expression of ccnd1 in midbrain cross-sections of

WT, znf644a morphants, and znf644b morphants (n=3 in each group). Red arrows emphasize mislocalized expression domains of ccnd1. (B) Immunostaining assays

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monitoring the activation of Caspase3 in WT, g9a morphant or znf644a morphant midbrain cross-sections as well as rescue experiments using co-injection of p53-MO at

72 hpf (n=3 in each group). Yellow arrows emphasize apoptotic cells. (C)

Immunostaining assays monitoring the expression of the cell proliferation marker PCNA in WT or znf644b morphant midbrain cross-sections (n=3 in each group). Yellow arrows emphasize proliferative cell populations.

3.5 Discussion

3.5.1 Neural-specific regulation of gene silencing during differentiation by G9a/ZNF644

The relatively tight neural progenitor cell-specific expression pattern of znf644a and znf644b as well as their overlapping functional requirements in regulating H3K9me2 in the promoter regions of progenitor-specific genes indicate that znf644a and znf644b serve as tissue-specific co-factors of G9a/GLP activity. Among the key features underlying a progenitor cell identity are (1) the expression of a unique complement of multipotency-related TFs and (2) the maintenance of a proliferative state. During the onset of differentiation, these attributes become suppressed, and RPCs shift into an amorphous stage wherein the transitioning cells either continue to transiently express multipotency-related TFs but no longer divide or, more rarely, no longer express multipotency-related genes but are still engaged in the cell cycle. Most of the literature on

RPCs indicates that multipotency-related TFs (i.e., Vsx2) function as upstream positive regulators of cell cycle progression that are critical to the maintenance of the proliferative state. For example, disruption of Vsx2 has been shown to lead to premature cell cycle

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exit as a downstream defect (Vitorino et al., 2009). While the maintenance of proliferation in early RPCs (during the expansive phase) requires the upstream activity of multipotency-related TFs, my data suggests that suppression of the proliferative multipotent state in the retina requires the activity of znf644a and znf644b.

While both znf644a and znf644b seemingly have overlapping functionality in terms of gene suppression via H3K9me2, the non-overlapping functional requirements vis-à-vis retinal development for znf644a and znf644b, as evident by distinctions in the characteristics of their morphant phenotypes (i.e., znf644b morphant retinas maintain a prolonged proliferative progenitor identity, whereas znf644a morphants fail to suppress progenitor-specific gene expression during differentiation) suggest multiple aspects of retinal differentiation are uniquely regulated by znf644a and znf644b. In the case of znf644b morphant retinas, there appears to be a tissue-specific developmental delay that caused that caused RPCs to maintain a proliferative progenitor cell fate for an extended period of time, resulting in the delayed emergence of differentiated retinal cells. In znf644a morphant retinas, there is persistent cellular proliferation that may cause a slight delay in differentiation, but, as a population, RPCs in these morphants are able to generate various types of differentiated neurons, albeit with impair cell viability, despite the persistent expression of certain RPC-specific genes. Hence, znf644a and znf644b have overlapping and non-overlapping regulatory roles in various aspects of RPC differentiation.

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3.5.2 Model for the role of the G9a-ZNF644 complex during RPC differentiation

Most of the literature on RPCs seems to indicate that multipotency-related TFs

(e.g., vsx2 and others) are upstream positive regulators of cell cycle progression genes. In addition, it is well established that proliferative and differentiating states are mutually exclusive; that is, proliferation/cell-cycle genes must be shut down for the occurrence of cellular differentiation. Using these principles as the backbone, I have derived a putative model outlining multiple roles for the G9a-ZNF644 complex during retinal differentiation

(Figure 3.20A). In this model, the suppression of G9a/H3K9me2 target genes, such as vsx2 and ccnd1, retinal cell viability, and the appropriate timing at which retinal differentiation commences are dependent on physical interactions between G9a and

ZNF644, which, in the case of zebrafish, consist of both overlapping and non- overlapping functions of two paralogues, znf644a and znf644b. (Figure 3.20A). In zebrafish, both znf644a and znf644b suppress the expression of multipotency-related TFs

(i.e., vsx2 and ccnd1), yet znf644b is uniquely responsible for ensuring that RPCs suppress their proliferative, multipotent identity and engage in cellular differentiation. I have provided evidence that the functions of both znf644 paralogues are critically dependent on physical association with g9a using “genetic cooperativity” assays as well as rescue experiments with mutant versions of ZNF644 that lack G9a/GLP binding capabilities.

While the model in Figure 3.20 was derived using zebrafish embryos as a model system, it can nonetheless be used to predict the regulatory roles of the G9a-ZNF644 complex in the developing mammalian retina. This extrapolation is supported by the

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demonstrated functional conservation of human ZNF644 with its zebrafish counterparts – recall that co-injection of human ZNF644 efficiently rescues the znf644a and znf644b morphant phenotypes. In this case, multiple regulatory aspects that are essential for retinal differentiation (i.e., the many g9a, znf644a and znf644b retinal phenotypes) are intimately coupled at the molecular level by the presence of a single, common G9a-

ZNF644 regulatory complex in the mammalian retina (Figure 3.20B). This possibility suggests highly complex and dynamic regulatory roles for the G9a-ZNF644 physical interaction in the developing CNS.

Figure 3.20: Model illustrating the roles of the G9a-ZNF644 physical association in retinal differentiation. (A) Model outlining the multiple aspects of retinal cell differentiation regulated by either/or the physical interactions between g9a-znf644a and g9a-znf644b based on the data presented in this thesis. (B) Predicted role of the mammalian G9a-ZNF644 physical interaction during retinal differentiation.

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At the level of gene regulation, it appears that both zebrafish znf644 paralogues repress the expression of similar set of G9a/H3K9me2 targeted genes in RPCs (Katoh et al., 2012). Indeed, I have demonstrated that two such targets, vsx2 and ccnd1, exhibit mislocalized expression throughout the retinas of znf644a and znf644b morphants. In addition, the mislocalized expression corresponds to – and is most likely caused by – the failed generation of H3K9me2 in the promoter regions of these two genes. Hence, because of the overlapping requirement of znf644a and znf644b in generating H3K9me2 in the promoter regions of certain RPC-specific genes, the key difference in paralogue function appears to be their requirement vis-à-vis differentiation: znf644a morphant retinas fail to maintain a proliferative progenitor identity (despite the persist expression of vsx2 and ccnd1) and undergo differentiation, whereas znf644b morphant retinas maintain a proliferative multipotent identity for an extended period of time before undergoing differentiation. In znf644b morphant retinas, progenitor cells appear to advance much more slowly in their development, resulting in a hypocellular proliferative retina that lacks apoptotic cells. Why znf644b, but not znf644a, exhibits such a regulatory function requires further investigation to determine whether subtle changes in their expression timing or whether there are paralogue-specific biochemical characteristics that impart unique functionality. In any case, the data collectively asserts both overlapping and non- overlapping roles for znf644a and znf644b in retinal differentiation – specifically, determining the onset of differentiation (znf644b morphant phenotype) and suppressing the expression of multipotency- and cell cycle-related genes (znf644a and znf644b morphant phenotypes).

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3.5.3 H3K9me2 marks differentiated neuronal cells in the retina

Large H3K9me2-marked chromatin territories, which are mostly absent in progenitor cells, are thought to emerge during lineage specification to play structural roles in maintaining a stable silenced epigenome (Wen et al., 2009; Chen et al., 2012). I observed H3K9me2-positive nuclei in differentiated neurons, which, I reason, likely represent expanded regions of H3K9me2-marked chromatin in these cells. The emergence of H3K9me2-positive nuclei was preceded temporally by cell cycle exit and the expression of lineage commitment factors (i.e., pax6a in amacrine cells, isl2b in

RGCs, and Zpr1 in cone photoreceptors) in newly born neuronal subtypes, suggesting the formation of expanded H3K9me2-marked chromatin may be a late-phase process that occurs within fully differentiated neuronal cells in vivo. Indeed, the prolonged proliferative RPC identity and delayed differentiation seen in znf644b morphant retinas led to a downstream absence in H3K9me2-positive nuclei at 72 hpf (Figure 3.16), perhaps indicating a further delay in the formation of such H3K9me2 marked nuclei.

Thus, znf644a and znf644b may facilitate the formation of H3K9me2 marked chromatin at isolated genomic regions (i.e., the promoters of vsx2 and ccnd1) in RPCs, and, then later in development, H3K9me2-marked chromatin may undergo progressive expansion as retinal cells differnetiate, culminating in H3K9me2-positive nuclei visible by microscopy.

3.5.4 Common cellular dependencies for G9a and ZNF644 in retinal and midbrain progenitors

In neural progenitors, ZNF644 appears to play a highly specialized role to impart target gene specificity for G9a-mediated silencing. The conserved atypical ZF motif of

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ZNF644, also found in WIZ, suggests a common vertebrate-specific innovation for modulating G9a function. Disruption of G9a or ZNF644 leads to defective RPC differentiation as well as seemingly analogous cellular effects in midbrain progenitors.

Despite distinct multipotency-related gene expression programs between the two neural substructures (i.e., vsx2 is uniquely expressed in RPCs), retinal and midbrain progenitors appear to have remarkably similar cellular dependencies on G9a and ZNF644, which may signify additional physical and functional interactions with progenitor subtype-specific cofactors or common target genes. Like WIZ in ES cell (Ueda et al., 2006), ZNF644 may serve as a molecular scaffold that facilitates the assembly of neural progenitor-specific repressor complexes at particular promoter or enhancer regions. Additional investigations are needed to characterize retina- and/or midbrain-expressed factors that may physically and/or functionally interact with G9a and ZNF644.

Roles for G9a in the timing of early lineage commitment and differentiation have been demonstrated in hematopoietic stem and progenitor cells (HSPCs) (Chen et al.,

2012) and ES cells (Yamamizu et al., 2012). I have provided evidence for analogous functions for a G9a-ZNF644 complex in neural progenitors, whereby retinal differentiation is delayed, seemingly due to a decreased rate of cell cycle progression, without subsequently affecting the birth order of newly formed neuronal subtypes.

Moreover, the Tropepe lab has demonstrated that blocking cell cycle exit can also lead to a lengthening of the RPC cell cycle leading to deficits in differentiation without concomitant apoptosis (Wong et al., 2010). As ZNF644 function seems to critically depend on its physical association with G9a, the intricate cellular defects underlying neuronal impairment seen in patients with 9q34 subtelomeric deletion syndrome may

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include, at least in part, defects in ZNF644 function. I also note that high expression of

ZNF644 has also been reported in the dentate gyrus of the adult mouse hippocampus

(Allen Brain Institute, Seattle, WA), where the proper timing of neural differentiation is important for neural circuitry integration (Farioli-Vecchioli et al., 2008). Further investigations are needed to determine possible functions for ZNF644 during adult neurogenesis.

3.5.5 Possible insights into high-grade myopia

It is generally thought that longitudinal stretching (or lack thereof) of the eye can alter axial elongation, resulting in myopia or hyperopia (Wallman and Winawer, 2004).

However, there are reports implicating developmental-related genes and other signaling pathways that regulate postnatal retinal growth in these disorders (Sundin et al.,

2005; Hysi et al., 2010; Fischer et al., 2006; Tkatchenko et al., 2006). Indeed, it has been demonstrated in primates that disruption of the CMZ, which harbors a proliferating RPC niche, can cause high-grade myopia (Tkatchenko et al., 2006). It is not clear, however, how the seemingly autosomal dominant SNPs reported (Shi et al., 2011; Tran-Viet et al.,

2012), which are situated both in the coding sequence and 3’UTR (and perhaps non-exon regions), affect ZNF644 function, whether it be at the level of altered mRNA stability, processing, or cognate protein levels and/or protein function.

Because high-grade myopia is typically a late onset disease, mutations in ZNF644 that segregate with the disease state may confer subtle functional deficiencies in the formation or activity of G9a-ZNF644 complexes that perhaps accrue over an extended period of time. Further investigations are needed to determine the roles of ZNF644 (and

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mutations associated with high-grade myopia) in controlling G9a activity postnatal retinal growth or differentiation in adult retinal stem cell populations.

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Chapter 4 Thesis summary and future directions

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4 Thesis summary and future directions 4.1 Thesis summary

In this thesis, I described my work investigating the composition of protein complexes associated with the histone methylation-related machinery. At the start of my work, most of our understanding about the composition, function and regulation of these complexes was limited to a handful of studies. To address this gap in understanding, I applied a systematic AP-MS approach using a recently developed lentiviral-based approach developed by our group (Mak et al., 2010; Ni et al., 2011). This body of work ties into a larger collaborative effort with the Greenblatt and Wodak labs aimed at mapping physical associations for various human chromatin-related protein complexes.

My contribution to this project was the systematic analysis of histone methylation-related complexes, as well as method development for the generic mass spectrometry and data analysis approach used for the project. A large dataset derived from this project has recently been submitted for publication.

As described in Chapter 2, I focused my own Thesis efforts on the biochemical characterization of histone methylation-related protein complexes of particular interest to me. As positive controls, I used several transcription-related complexes (RNAPII,

Mediator and CPSF complexes) to optimize all the pipeline (i.e., purification, LC/MS/MS and PPI scoring) procedures. I applied previously established AP-MS data analysis tools, particularly the SAINT algorithm developed in part by Dr. Gingras in our Department

(Choi et al., 2011). Applying a stringent benchmarking and filtering approach allowed me to generate a high-confidence interaction network comprising 573 PPIs, of which only roughly half (312) were previously annotated in a public interaction database (Turner et

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al., 2010; Chatr-Aryamontri et al., 2013). While there are relevant unannotated interactions reported in the literature, I documented several putatively novel interacting protein subunits of especially noteworthy interest. In particular, I first focused on exploring the significance of a novel set of RNAPII-interacting protein, RPRD1A,

RPRD1B and RPRD2, which was surprising given that RNAPII had been so heavily studied to date and given how central this complex is to transcriptional control in general, and then more extensively to characterizing the functional significance of the association a between the uncharacterized protein ZNF644 and the H3K9 HMT G9a, given how likely I anticipated this unexpected complex was likely to play in the epigenetic regulation of developmental expression programs, which was in fact borne out in my follow up mechanistic studies.

The uncharacterized RNAPII co-factors RPRD1A, RPRD1B and RPRD2 proteins each contained N-terminal CTD-interacting domains (CIDs), which are also situated in certain yeast proteins involved in transcriptional termination (Rtt103, Pcf11 and Nrd1). Reciprocal AP-MS experiments using RPRD1A and RPRD1B as “baits” revealed that these proteins form an RNAPII-containing complex that also contains known negative regulators of transcription (GRINL1A and RECQL5) as well as the putative CTD-S5 phosphatase RPAP2, of which the putative yeast orthologue (Rtr1) has been reported to regulate a shift in CTD phosphorylation, from S5P during transcriptional activation to S2P during transcriptional elongation (Mosley et al., 2009). Since the CIDs in Rtt103, Pcf11 and Nrd1 are known to bind phosphorylated CTD repeats (Vasiljeva et al., 2008; Lunde et al., 2010; Meinhart and Cramer, 2004), I performed co- immunoprecipitation experiments using monoclonal antibodies that recognize specific

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CTD phosphorylation isoforms (i.e., S2P, S5P and S7P) and lysates from cells stably expressing VA-tagged RPRD1A or RPRD1B. This analysis revealed that the RPRD proteins preferentially associate with phosphorylated forms of RNAPII. This work was coupled with subsequent additional RPRD-related drill down studies performed by Dr.

Zuyao Ni, a postdoctoral fellow in the Greenblatt lab, and we published our results together as co-primary authors in the journal Transcription (Ni et al., 2011).

My AP-MS experiments with tagged HMT G9a indicated that it stably co-purified with, as expected, its paralogue and known heterodimerization partner GLP, (Tachibana et al., 2005) and WIZ, which had previously been identified as a G9a/GLP-interacting protein in ES cells (Ueda et al., 2006), and a second, but less characterized, large C2H2

ZF motif-containing protein, ZNF644. Reciprocal AP-MS experiments using GFP-tagged

ZNF644 confirmed the interaction with G9a and GLP, but also, somewhat unexpectedly, that ZNF644 and WIZ can reside in the same G9a/GLP-containing complex. In addition,

I determined that an atypical ZF motif situated in the C-terminal region of ZNF644 directly binds to the G9a SET domain, as is the case for WIZ (Ueda et al., 2006), and is both necessary and sufficient for the physical association with G9a, GLP and WIZ.

Moreover, I used lentiviral-based shRNA-mediated knockdown in cell culture to demonstrate that, like G9a, ZNF644 regulates global levels of H3K9me2, indicating that

ZNF644 plays a likely plays a critical role in modulating the enzymatic activity and/or chromatin targeting of G9a/GLP.

Given that a rapidly expanding body of literature now points to critical roles for

G9a/GLP in diverse physiological contexts, I was determined to figure out the biological role of the G9a-ZNF644 interaction in vivo. At the time I embarked on these studies, a

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new paper was published demonstrating that mutations in the ZNF644 link segregated with autosomal dominant inherited forms of high-grade myopia (Tachibana, 2005).

Hence, I was particularly intrigued by the possibility of determining how G9a and

ZNF644 might cooperatively function within the context of the retina. Incidentally, it had recently been demonstrated that zebrafish G9a functioned cooperatively with DNMT3 during embryogenesis (Rai et al., 2010). Among the defects they observed in g9a morphant embryos were defects in neurogenesis, both in the retina and in the brain.

Accordingly, I set up and pursued a collaborative effort with members of the Tropepe lab, who are experts in zebrafish neurobiology, and worked particularly close with his graduate student Loksum Wong (recently awarded a PhD for independent work).

Chapter 3 of my thesis describes our subsequent investigations and key findings, which together with my work in Chapter 2 forms the basis for a co-first authored manuscript that is currently under review for publication.

As is commonly observed in zebrafish, the mammalian ZNF644 gene has two zebrafish paralogues, which I termed znf644a and znf644b. Each paralogue has selectively retained a reduced complement of C2H2 ZF motifs (i.e., human ZNF644 contains 7 C2H2 ZF motifs, whereas Znf644a and ZNF644 each contain 5), suggesting possible functional divergence (either full or partial). However, both znf644 paralogues contain atypical C2H2 ZF motifs (i.e., the G9a-interacting module) in their C-terminal regions, suggesting that at least their physical association with G9a is likewise conserved.

I also found that, in contrast to the broad expression of G9a during zebrafish embryogenesis (Rai et al., 2010), the expression patterns of both znf644 paralogues were both temporally and spatially regulated and largely overlapping. Specifically, both

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paralogues were transiently expressed at the 24 hpf time point, a stage at which proliferating neural progenitor are preparing withdraw from the cell cycle and differentiate. Hence, I used a morpholino (MO)-based knockdown approach to determine how G9a and ZNF644 might co-regulate the development of these regions, with particular emphasis on the retina, given the association of ZNF644 with high-grade myopia.

Genetic characterization of the neuronal/retinal functions of G9a and two ZNF644 paralogue genes in zebrafish turned out to be a challenging and complex process, as individual disruption of these genes gave rise to non-overlapping phenotypes. While characterizing the roles of g9a and the znf644 paralogues in the zebrafish retina, Katoh and colleagues (Katoh et al., 2012) published their findings regarding a role for G9a in mouse RPCs in suppressing apoptosis and suppressing the expression of a set of multipotency- and cell cycle-related genes. Among the genes silenced by G9a-mediated

H3K9me2 were the multipotency-related TFs Vsx2, Lhx2 and Hes1, of which Vsx2 has a well-characterized orthologue with analogous function in the retina (Vitorino et al.,

2009), as well as several types of cyclins, such as CyclinD1/ccnd1, that drive cell cycle progression.

My data implicates G9a as the primary HMT responsible for the suppression of progenitor-specific genes involved in maintaining a multipotent proliferative state. As observed in mouse RPCs (Katoh et al., 2012), the RPCs of g9a morphant zebrafish retinas were able to withdraw from the cell cycle, but were highly apoptotic. These morphant retinas also failed to give rise to virtually any type of differentiated neuron. In contrast, I obtained evidence that, as a population, the RPCs of znf644a morphant retinas

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(which resulted in a near complete loss of mature znf644a mRNA) were able to withdraw from the cell cycle and give rise to various types of differentiated retinal neurons.

However, these differentiated neurons exhibited persistent expression of multipotency- related (i.e., vsx2) and cell cycle-related (i.e., ccnd1) and, consequently, underwent apoptosis. The RPCs of znf644b morphant retinas likewise showed persistent expression of progenitor-specific genes (i.e., vsx2 and ccnd1), but these cells remained in a proliferative state for a prolonged period of time, resulting in temporally delayed differentiation; nevertheless, these RPCs eventually withdrew from the cell cycle and eventually gave rise to a seemingly normal complement of retinal neurons.

I applied a “genetic cooperativity” approach to better tease out the possible functional dependencies between g9a and the two znf644 paralogues. Commonly used in zebrafish genetics, this technique involves the co-injection of MO doses that individually do not give rise to a detectable phenotype and indeed had been used previously to reveal functional cooperativity between G9a and DNMT3 (Rai et al., 2010). As anticipated, I found compelling evidence of functional cooperativity between all three proteins.

Combining subphenotypic doses of MOs targeted against G9a and Znf644a gave rise to an apoptotic retina reminiscent of the higher MO dose g9a morphant retina, whereas combining subphenotypic MO doses of G9a and znf644b, gave rise to a prolonged proliferative retina reminiscent of higher dose znf644b morphant retina. In fact, there was also functional cooperatively among the znf644 paralogues, as co-injection of subphenotypic doses of MOs targeted against znf644a and znf644b gave rise to a prolonged proliferative retina, again as observed in znf644b morphants.

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Of particular note, these assays appear to have revealed some interesting insights regarding the relationship between the expression of multipotency-related TFs and cell cycle progression in RPCs during differentiation. Most of the literature on RPCs supports the paradigm that multipotency-related TFs (vsx2 and ccnd1) are critical upstream positive regulators of cell cycle progression genes. The evidence for this is primarily based in genetic studies where perturbation (e.g. mutation) of multipotency-related TFs in human, mouse and zebrafish causes premature cell cycle withdrawal, differentiation, and microphthalmia. Whether this dependency is maintained throughout the full development of RPCs is currently unknown, but it has been widely assumed that retinal differentiation likely involves a “domino effect” where the suppressed expression of multipotency- related TFs alone results in downstream withdrawal from the cell cycle. If this were indeed the case, the mislocalized expression of vsx2 and ccnd1 would have been sufficient to cause znf644a morphant retinas to maintain a progenitor identity for an extended period of time rather than undergoing differentiation. Accordingly, by manipulating the g9a-znf644a and g9a-znf644b complexes via MO treatments, I have been able to generate evidence that brings into question whether this rigid hierarchy holds true in late RPCs prior to differentiation. Specifically, I observed both RPCs lacking the detectable expression of multipotency-TFs (i.e., vsx2) were nonetheless able to maintain a proliferative state, as well as RPCs that failed to suppress the expression of multipotency-related TFs (i.e., vsx2) and yet withdrew from the cell cycle and terminally differentiated anyways. These findings argue against the “domino” model and suggest a more involved suppressive mechanism during retinal differentiation (see Figure 3.20), wherein the proper timing and rate of retinal development and the suppression of an RPC

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gene expression program must be coupled at the molecular level (and hence temporally linked) by the G9a-ZNF644 physical association. Collectively, my body of work provides the first detailed mechanistic characterization of the role of ZNF644 development, linking this its function to G9a/H3K9me2-mediated regulation of gene expression during neuronal differentiation in both the retina and the midbrain.

4.2 Future directions

4.2.1 Characterization of novel components of histone methylation-related complexes

The histone methylation-related PPI network has revealed several novel putative components including unexpected regulators of histone methylation. Much like I demonstrated the role of ZNF644, each of these novel candidates holds the possibility of shedding insight regarding how histone methylation and the chromatin-modifying machinery function in physiological contexts. Much structure-function analysis remains to be done to determine the regulatory significance of other promising novel interacting proteins, such as BOD1L that co-purified with the COMPASS-like complexes, and

C10orf12 with PRC2, just as we have attempted to glean for the RPRD proteins found with RNAPII.

4.2.1.1 Regulation of G9a/GLP by ZNF644 and WIZ

I spent considerable time during my PhD studies to investigate the function of

ZNF644 vis-à-vis G9a/GLP. However, it is still unclear how ZNF644 regulates the function of G9a and GLP. One possibility is that, via its array of canonical ZN fingers, it serves as a sequence-specific nucleic acid targeting factor. With the Hughes lab, I assayed the 5 C2H2 ZF cluster in the central portion of human ZNF644, as well as the

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two C2H2 ZF clusters in the zebrafish versions of ZNF644, for DNA binding activity in vitro using protein binding microarrays (Bulyk, 2007). However, I failed to detect any sequence-specific binding in the assay (data not shown). Likewise, ChIP-seq experiments from HEK293 cells expressing GFP-tagged full-length wild-type ZNF644 failed to reveal significant DNA binding (data not shown). In the case of ChIP-seq, it may be that

ZNF644 only binds DNA at specific phase of the cell cycle, perhaps during S-phase when G9a/GLP are thought to function at site of DNA replication to “copy” their methylation mark to newly formed nucleosomes (Esteve et al., 2006). Moreover, ZNF644 may not directly bridge G9a/GLP to specific genomic regions through site-specific DNA binding. At present, I prefer the possibility that ZNF644 regulates the formation of a neural progenitor-specific G9a/GLP repressor complex, serving to scaffold the HMTs to other TFs and co-repressor machinery, much in the same way WIZ seems to serve as a scaffold molecule that bridges G9a and GLP to the CtBP co-repressor machinery in ES cells (Ueda et al., 2006). In addition, the fact that ZNF644 co-purified with WIZ begs the question as to the relationship between these two similar proteins. What are the differences between WIZ and ZNF644 and how do they differentially regulate G9a/GLP function? Other than limited data in ES cells, the tissue expression and developmental contexts in which WIZ might regulate the function of G9a/GLP is currently unknown.

However, based on unpublished human tissue proteomics data from a collaborating group, it appears that G9a, GLP, WIZ and ZNF644 may be co-expressed in several types of adult cells of the neural, germ-line, and hematopoietic systems (data not shown). It will be interesting, then, to see whether the proposed new roles of the G9a-ZNF644

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complex I have report in the developing CNS has commonality with other physiological systems.

4.2.2 Further tissue-specific characterization of HMT protein complex function

Protein interactions are often dynamic (i.e., transient), and the composition of complexes may change throughout cellular processes such as the cell cycle, as well as in response to external stimuli. In this study, I generated a static snapshot of the histone methylation-related machinery from proliferating cells in culture. However, the diverse tissue-specific roles of histone methylation-related proteins and their apparent role in various disease contexts likely involve condition-specific regulatory cofactors. This has already been demonstrated in certain physiological contexts. For example, the microcephaly-linked protein ZNF335 has recently been demonstrated to be a regulatory component of a vertebrate-specific COMPASS-like complex that regulates the expression of essential neural-specific master regulators, such as NRSF/REST (Yang et al., 2012).

4.3 Integration of HMT complex function with master transcriptional regulatory networks and upstream signalling pathways

While the histone methylation and associated chromatin-modifying machinery are critical for controlling DNA templated processes and chromatin structure, these protein complexes are part of a highly integrated system that also involves inputs from upstream cell signalling pathways as well as transcriptional regulatory networks by DNA-binding

TFs. As described earlier, the identity of a cell – which includes its development, biochemical behavior, and physiological responses to environment cues – is largely determined by the set of TFs that it expresses. This is best represented by induced

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pluripotency, which now forms the backbone of modern efforts in the field of regenerative medicine. However, in addition to their DNA binding domains, TFs also contain effector domains that enable them to both interact physically with each other, allowing for greater specificity via combinatorial regulation, and with various components of the basal transcriptional machinery as well as various chromatin modifying and remodeling enzymatic complexes to the promoter they regulate (Frietze and Farnham, 2011). Moreover, activation (and inactivation) of TF function can often be regulated by upstream membrane and cytosolic signalling pathways, via modulations in

PTM, proteolytic processing, and/or nuclear localization (to name a few).

In the context of the research I presented here, it is unclear how the activities of the HMT protein and complexes I investigated are dynamically regulated in vivo and how these operate within the broader context of integrative biological systems. For example, it remains to be determined what drives the seemingly transient and tissue- specific expression of ZNF644 in retinal and midbrain progenitor cells. Is it tied into other master neurogenic signalling pathways that stimulate cell cycle withdrawal and differentiation in the retina, such as Shh and FGF (Agathocleous and Harris, 2009), or might it be related to common cellular processes, such as cell cycle rate or specific types of cell-cell contacts/adhesion?

4.3.1 Determining the molecular factors regulating the target gene specificity of histone methylation-related complexes

Revealing the precise mechanisms whereby a transcriptional and/or chromatin- modifying complex is recruited to, and silences, target genes are often a complicated step-wise process (Voss and Hager, 2013). As I described earlier, it is known that the

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recruitment of a G9a/GLP-containing complex to the Oct4 locus during ES cell differentiation involves concomitant histone deacetylation and DNA methylation, yet despite candidates, the exact molecular factors that associate first with the Oct4 locus and in turn recruit repressors like G9a are still incompletely resolved. Of particular interest are non-coding RNAs and their potential contribution in determining the target gene specificity of histone methylation-related complexes. For example, long intragenic ncRNAs (lincRNAs) are known to directly interact with components of HMT complexes and appear to drive them to specific loci. Perhaps the best-characterized example of this targeting is the recruitment of PRC2 to the inactive X chromosome via specific recognition of stem-loop structures situated in the lincRNA Xist by the SUZ12 subunit

(Kanhere et al., 2010). The building of a genome-wide RNA-chromatin protein interaction network is critical for understanding ncRNA function vis-à-vis epigenetic gene regulation, followed by detailed molecular investigation into how these complexes are assembled and regulated.

4.3.2 How are histone methylation-related complexes related to disease processes? In addition to understanding the cellular and molecular underpinnings of biological systems and the role of epigenetics in development, understanding how biochemical regulation by protein complexes can often lead to insights regarding aberrant cellular processes that initiate and propagate disease processes. There is now compelling evidence that HMTs affect transcription in disparate manners depending on cellular context. For example, in castration-resistant prostate cancer cells, the core enzymatic component EZH2 can reside in either a conventional PRC2 complex that exerts H3K27 methyltransferase ability as well as a separate, independent transcriptional activation

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complex bound by the androgen receptor that drives an oncogenic gene expression program (Xu et al., 2012). Because the histone methylation-related protein machinery plays critical roles in shaping the landscape of the mammalian epigenome in diverse cellular and physiological contexts, continuing efforts to profile chromatin modification marks and complex recruitment during normal developmental and disease states should continue to be informative. It is anticipated that such efforts will aid in revealing the mechanistic underpinnings of multicellular development, cell fate decisions, and disease processes. Given the importance of histone methylation-related and other types of chromatin-modifying complexes as an emerging class of therapeutic importance

(Copeland, Moyer and Richon, 2013; Copeland, Solomon and Richon, 2009; Arrowsmith et al., 2012), continual research conducted along these lines may very well even form the basis of future directed clinical strategies.

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