Temporal Regulation of Chromatin Organization During C. Elegans Embryogenesis

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Temporal regulation of chromatin organization during C. elegans embryogenesis

A dissertation presented

Beste Mutlu

The Department of Molecular and Cellular Biology

in partial fulfillment of the requirements

for the degree of

Doctor of Philosophy

in the subject of

Biochemistry

Harvard University

Cambridge, Massachusetts

April 2018

Dissertation Advisor: Professor Susan E. Mango Beste Mutlu

Temporal regulation of chromatin organization during C. elegans embryogenesis

Abstract

In all eukaryotic cells, the genetic material is organized into a complex structure composed of DNA and protein: chromatin. Organization of chromatin structure in the nucleus influences all functions of the genome including gene expression and is dynamically regulated based on the needs of the cell. Chromatin re-organization is critical for avoiding the development and progression of pathological situations.

In most animals, chromatin undergoes major restructuring as embryos develop.

Accessible chromatin is a feature of naïve embryonic cells and is lost as cells become more specialized. As cells mature, the nucleus shrinks, chromatin compacts and silent heterochromatin domains are generated. Heterochromatin is typically associated with repressive post-translational modifications such as histone H3 lysine 9 (H3K9) methylation. While the machinery that organizes chromatin is known, the molecular cues that initiate chromatin re-organization in the early embryo are elusive.

Using C. elegans as a model, I found that a conserved H3K9 methyltransferase is critical for the onset of heterochromatin formation. H3K9 methylation by MET-2/SETDB1 is temporally regulated by two conserved factors identified in this study. Nuclear accumulation of MET-2 and its binding partners are rate-limiting for heterochromatin

iii formation. Polymerase II transcription, RNA interference pathways that are known to assemble heterochromatin, or mechanisms that rely on cell counting in the embryo do not dictate the timing of this transition. Instead, slowing the early embryonic cell cycles leads to precocious heterochromatin, suggesting that absolute time after fertilization is a key determinant. My studies delineate the temporal regulation of MET-2 and its binding partners, and their role in chromatin re-organization during embryogenesis. Orthologs of these worm proteins likely have similar roles in re-organizing chromatin in other systems and are implicated in human disease.

iv TABLE OF CONTENTS

ABSTRACT……………………………………………………………………………………...iii

ACKNOWLEDGEMENTS…………………………………………………………..………...vii

CHAPTERS

1. INTRODUCTION………………………………………………………………………..1

1.1 Overview……………………………………………………………………………..2 1.2 Chromatin re-organization during development…………………………………3 1.3 SETDB1 mediated H3K9 methylation…………………………………………..19 1.4 C. elegans embryogenesis as a model for chromatin re-organization………21 1.5 References…………………………………………………………………………25

2. REGULATION OF A HISTONE METHYLTRANSFERASE TIMES THE ONSET OF HETEROCHROMATIN FORMATION IN C. ELEGANS EMBRYOS………...49

2.1 Abstract…………………………………………………………………………….50 2.2 Main Text…………………………………………………………………………..51 2.3 Materials and Methods……………………………………………………………65 2.4 References…………………………………………………………………………82

3. THE ONSET OF HETEROCHROMATIN FORMATION IS DICTATED BY TIME AFTER FERTILIZATION………………………………………...... 88

3.1 Abstract…………………………………………………………………………….89 3.2 Introduction………………………………………………………………………...90 3.3 Results…………………………………………………………..………………….92 3.4 Discussion………………………………………………………………………..106 3.5 Materials and Methods………………………………………………………….110 3.6 References…………………………………………………………………….…114

v 4. CONCLUSION………………………………………………………………………..122

4.1 Overview……………………………………………………………………..…...123 4.2 Summary and Discussion..……………………………………………………..123 4.3 Future Directions…………………………………………………………………128 4.4 Concluding Remarks…………………………………………………………….134 4.5 References……………………………………………………………………….135

APPENDIX……………………………………………………………………………………143 Supplementary Figures for Chapter 2……………………………………………...144 Table A.1 List of MET-2::GFP interactors…………………………………………151 Table A.2 List of 3xFLAG::MET-2 interactors…………………………………..…153 Experimental differences for MET-2::GFP and 3xFLAG::MET-2 IP…………....160 Conservation of MET-2 in other organisms……………………………………….161 Analysis of MET-2::GFP hubs in lin-65 and arle-14 mutants……………………162 Analysis of LIN-65 protein in met-2 and arle-14 mutants………………………..164 References……………………………………………………………………………165

vi Acknowledgements

I would like to thank:

• My wonderful family for their unwavering support and encouragement.

• My supervisor Dr. Susan E. Mango for guiding and mentoring me throughout my

PhD, for inspiring me to stand up for my opinions and most importantly for

reminding me to enjoy the journey.

• Dr. Alex Schier and Dr. Danesh Moazed for extremely valuable feedback about

my project and manuscripts, and for always taking the time to help.

• Dr. Vlad Denic for experimental help with biochemistry protocols and intellectual

discussions at committee meetings.

• My collaborators Dr. David E. Hall, Dr. John Yates III, Dr. James Moresco, Claire

Reardon and Dr. John Gaspar.

• Former and current members of the Mango Lab for creating a fun and collaborative

work environment and listening to countless practice talks. Especially Dr. Stephen

Von Stetina and Dr. Huei-Mei Chen for their mentorship and friendship, and Sabine

Keppler-Ross for her help with worm injections.

• All the MCO administrative staff, especially Michael Lawrence, Patty Perez,

Debbie Maddelena, Allie Pagano, Katie Scrocca and Fanuel Muindi.

• American Association for University Women (AAUW) for providing me funding

through an International Fellowship.

vii

CHAPTER 1

INTRODUCTION

1.1 Overview

The primary role of the nucleus is storage of genetic information, which involves physical and functional compartmentalization of the genome. DNA wraps around histone octamers to form nucleosomes and chromatin fibers, which are spatially organized into topological domains, compartments and chromosome territories (Dixon, Gorkin, and Ren

2016). Depending on the needs of the cell, chromatin organization can be regulated locally at the level of individual nucleosomes or globally at the level of chromatin domains.

Chromatin can be broadly separated into two states based on transcriptional activity, sequence features, post-translational histone modifications and compaction.

Euchromatin is defined as genomic regions that are accessible and transcriptionally active, whereas heterochromatin is characterized by condensed chromatin, a high frequency of repetitive sequences and silencing post-translational histone modifications

(Rice et al. 2003; Rübe et al. 2011). The ratio of euchromatin and heterochromatin in the nucleus is not static (Fadloun, Eid, and Torres-Padilla 2013). Rather, chromatin is a dynamic structure that regulates access to DNA. Dynamic re-organization of chromatin is crucial for responding to stimuli and orchestrating cell-type specific developmental programs. For instance, embryos undergo large-scale chromatin re-organization during the first hours of development (Politz, Scalzo, and Groudine 2013).

In this chapter, I will first review current methodologies to study chromatin organization, the developmental changes that have been observed using these approaches and the function of chromatin re-organization during development. In the second part, I will focus on SETDB1 mediated H3K9 methylation and its regulation during

2 development. Last, I will present C. elegans embryogenesis as a model for studying large- scale chromatin re-organization during development and the role of H3K9 methylation in worms.

1.2. Chromatin re-organization during development

• Basics of chromatin organization

To fit the whole genome into the small volume of the nucleus, cells achieve 10,000-

20,000 fold compaction (Woodcock and Ghosh 2010), while allowing proteins involved in transcription, replication and repair to access DNA. Nucleosomes, building blocks of chromatin, constitute the first level of chromatin compaction and can be altered to modulate access to DNA. Nucleosomes consist of core histones (H2A, H2B, H3, H4), and

~150 bp of wrapped DNA (Luger et al. 1997), with H1 linker histones binding to the linker

DNA at the nucleosome entry and exit sites to stabilize the entire complex (Syed et al.

2010). There are three ways of regulating chromatin on the level of nucleosomes: i) Histone modifications: The variability in the composition of the nucleosome provides a molecular means by which the cell regulates local chromatin organization. A large number of residues on the unstructured N-terminal histone tail can be decorated with diverse post- translational modifications, including but not limited to methylation, acetylation and phosphorylation (Kouzarides 2007). Combinations of histone modifications at specific loci, or “chromatin states”, correlate with functional features such as promoters, enhancers, transcribed regions or silent domains and are largely conserved across species (Ho et al. 2014).

Histone acetylation was the first modification discovered with a role in transcriptional activation in the 1960s (Phillips 1963; Pogo, Allfrey, and Mirsky 1966). It is thought that acetylation regulates transcription in two ways. First, it neutralizes the positive charge of lysine residues on the histone tail, thus weakening the DNA- nucleosome interactions and rendering DNA more accessible to the transcription machinery (Dion et al. 2005). For instance, incorporation of acetylated Histone H4 Lysine

16 (H4K16ac) into nucleosome arrays inhibits the formation of compact chromatin fibers in vitro (Shogren-Knaak et al. 2006). Second, acetylated residues can be recognized by proteins that contain a bromodomain module (Zeng and Zhou 2002). Bromodomain factors Bdf1/2 associate with and recruit the general transcription factor complex TFIID to active genes in euchromatin (Matangkasombut et al. 2000; Durant and Pugh 2007).

Histone methylation does not directly affect the charge on the histone tail but is thought to have indirect effects through the recruitment of downstream effectors.

Methylated lysines can be bound by many domains, including Tudor, chromo and MBT

(Taverna et al. 2007). Heterochromatin Protein 1 (HP1), a transcriptional repressor and building block of heterochromatin, binds methylated Histone H3 Lysine 9 (H3K9me) with its chromodomain (Lachner et al. 2001). H3K9 and H3 Lysine 27 (H3K27) methylation are typically associated with gene silencing, chromatin compaction and heterochromatin

(Rice et al. 2003; Francis, Kingston, and Woodcock 2004). Overall, post-translational histone modifications are extremely diverse, and a subset of these marks have well studied roles in regulating transcription, replication, and repair (Table 1.1).

Table 1.1. A short list of well-known histone modifications and their function, reviewed in (Kouzarides 2007). Histone Site Modification Enrichment Function H3 K4 methylation Active gene promoters Transcription activation H3 K36 methylation Active gene bodies Transcription elongation H3 K9 methylation Repeats, silent genes Transcription silencing, DNA repair H3 K27 methylation Silent genes Transcription silencing H4 K20 methylation Gene bodies Gene expression, DNA replication and repair H3 K27 acetylation Enhancers Gene activation H3 K56 acetylation Transcription start sites Gene activation, DNA repair ii) Histone variants: Histone variants are a second way of altering the composition of the nucleosome. They are encoded by distinct genes that have different nucleotide sequences compared to their major counterparts and can be substituted into the nucleosome core at specific genomic loci (Buschbeck and Hake 2017). For instance, acetylated variant H2A.Z is enriched at active promoters and enhancers (Jin et al. 2009), whereas hypo-acetylated H2A.Z accumulates in heterochromatic regions (Hardy et al.

2009). Both histone modifications and variants affect nucleosome stability and dynamics, altering the local fabric of the chromatin fiber and influencing gene expression (Luger,

Dechassa, and Tremethick 2012). iii) Nucleosome remodeling: Nucleosome positioning throughout the genome determines accessibility of binding sites to transcription factors and transcriptional machinery, and thus affects processes such as transcription, DNA repair, replication (Radman-Livaja and

Rando 2010). Movement or exchange of nucleosomes by ATP dependent chromatin

5 remodelers regulates nucleosome occupancy and promoter architecture (Hargreaves and Crabtree 2011). There is competition between nucleosomes and transcription factors, so remodeling factors can deposit or clear repressive nucleosomes from transcription start sites to regulate gene expression (Cairns 2009). Nucleosome depleted regions typically represent active genes in euchromatin (Lee et al. 2004).

• Different methods to study chromatin organization

The historical basis for the distinction between euchromatin and heterochromatin is the differential staining properties observed by Heitz (Heitz 1928). Modern assays that analyze different aspects of chromatin organization corroborate the segregation of the genome into euchromatin and heterochromatin domains, but they also paint a more accurate and complex picture about chromatin states. While each assay has its advantages and caveats, together, they provide a powerful tool kit to study chromatin organization.

A) Elucidating genome-wide distribution of histone marks, variants and associated chromatin factors

Large scale mapping of histone modifications and related structures has become a powerful tool for characterizing chromatin organization (Brown and Celniker 2015).

Chromatin immunoprecipitation combined with sequencing technologies (ChIP-Seq) provides information about the genomic distribution of these marks and proteins associated with them. Integration of these maps with RNA expression data has revealed the role of chromatin organization in regulating transcription (summarized in Table 1.1) and moved the field from a gene-centric view to genome-wide scale (Zhou, Goren, and

Bernstein 2011). While these results are informative, they rely on correlation to make inferences about function and focused mechanistic studies are still required.

ChIP-Seq is useful for studying distribution of marks, however comparison of global levels between data-sets can be difficult. The difference in levels depends on the computational analysis methods and the threshold used and can cause controversy

(Wen, Wu, and Shinkai 2009; Lienert et al. 2011).

An alternative method to study distribution and levels of histone marks is immunofluorescence (IF). IF can be used to study global changes in the levels of marks and analyze their spatial distribution in the nucleus (e.g. proximity to the nuclear periphery, diffuse vs. punctate staining pattern) (Yuzyuk et al. 2009; Towbin et al. 2012).

With both techniques, close attention must be paid to antibody specificity.

B) Analyzing differential sensitivity of nucleosome occupied vs. depleted regions

Chromatin accessibility can be measured by utilizing three key methods that rely on differential sensitivity of nucleosome depleted vs. occupied chromatin to i) cleavage by non-specific nucleases such as micrococcal nuclease (MNase) and deoxyribonuclease (DNAse), ii) breakage by sonication and iii) integration of transposons

(Zentner and Henikoff 2014). Nucleosome depleted chromatin is more sensitive to all three perturbations. Combination of these methods with high-throughput sequencing technologies have enabled genome-wide mapping of nucleosome occupancy by nuclease based MNAse-Seq (Ercan et al. 2011), sonication based FAIRE-Seq (Giresi and Lieb 2009) and transposase based ATAC-seq (Gangadharan et al. 2010).

C) Measuring mobility of histones and chromatin binding proteins

The rate at which chromatin binding proteins associate with DNA is thought to influence gene expression (Voss and Hager 2008). Most transcription factors are highly mobile and transiently interact with chromatin in vivo, their residence times on chromatin ranging between 5-25 seconds (Phair et al. 2004). Core histones are relatively stable and their residence time is several hours, whereas that of a H1 linker histone is estimated to be 3 minutes (Catez, Ueda, and Bustin 2006). Studies have shown that the kinetics of exchange for histones and chromatin factors is affected by the condensation level of chromatin (Meshorer et al. 2006; Bošković et al. 2014; Cheutin et al. 2003).

Fluorescence Recovery After Photobleaching (FRAP) is the most commonly used technique used to track mobility of histones and chromatin binding proteins. In FRAP, a selectively photobleached region of the cell is monitored as fluorescence is re-established and the recovery process contains information about the mobility of proteins. Binding interactions with large, relatively immobile substrates such as chromatin alter the recovery rate, and can be used as a tool to quantify protein interactions with chromatin (Mueller et al. 2012).

D) Defining structural chromatin domains based on contact frequency

Chromatin conformation capture methods such as Hi-C can be used to study the

3D organization of the genome in the nucleus. Recent advances in these methods have identified a hierarchical series of structural domains in the nucleus (Figure 1.1), which point to a higher-order organization in 3D beyond nucleosomes (Kaiser and Semple

2017). These structures are physical entities defined by contact frequency: self- association and insulation from other domains (Lieberman-Aiden et al. 2009).

Figure 1.1. Layers of chromatin organization. DNA is wrapped around histones to form nucleosomes and chromatin fibers, which are spatially organized through looping interactions into Topologically Associated Domains (TADs), compartments and chromosome territories. TADs correlate with but do not rely on histone modifications or associated proteins.

For instance, each chromosome occupies a territory in the interphase nucleus.

Chromosome territories are divided into A and B compartments, which are 5-10 Mb large domains and strongly correlate with active and inactive chromatin, respectively (Sexton et al. 2012). Compartments are further divided into topologically associating domains

(TADs), ~1Mb regions with distinct boundaries containing loci that interact more with each

9 other than loci in different TADs (Dixon et al. 2012). In mammals, TADs contribute to establishing regulatory enhancer-promoter interactions through chromatin looping

(Tolhuis et al. 2002; Ptashne 1986), which typically occur within the same TAD

(Schoenfelder et al. 2015). TADs align with, but do not rely on epigenetic signatures such as histone modifications or associations with the nuclear lamina (Nora et al. 2012).

Instead, chromatin insulators such as CTCF or Cohesin have emerged as central components for defining TAD boundaries (Matharu and Ahanger 2015).

E) Identifying electron-lucent vs. electron-dense regions in the nucleus

Global segregation of the genome into euchromatin and heterochromatin can be detected by analyzing the ultrastructure of the nucleus using electron microscopy. These techniques detect heterochromatin as electron-dense regions (EDRs) amid electron lucent euchromatin. The designation of EDRs as heterochromatin rests on immunogold labeling experiments that show EDRs are enriched for methylated H3K9 and H1 linker histones, and lack modifications associated with active transcription such as acetylated

H4 (Studencka et al. 2012; Wirth et al. 2009; Cmarko et al. 2002; Rübe et al. 2011).

• Developmental changes in chromatin organization

Accessible chromatin is a feature of pluripotent cells and is lost as cells transit towards differentiation (Gaspar-Maia et al. 2011). Early mouse embryos, embryonic stem cells (ESCs) and planarian neoblasts all share large nuclei with a uniform distribution of euchromatin (Niwa 2007). As cells mature, nuclei shrink, chromatin compacts and the genome is broadly segregated into euchromatin and heterochromatin (Politz, Scalzo, and

Groudine 2013). Multiple assays have revealed significant changes in the chromatin landscape during differentiation (Table 1.2).

Table 1.2. Summary of assays used to study different aspects of chromatin organization during development. Aspect of chromatin Purpose of Assay Technique Developmental organization change Local and global Histone modifications, Elucidating genome- ChIP-Seq changes in the variants, associated wide distribution of Immunofluorescence distribution and proteins marks and proteins. levels of certain marks. Analyzing differential sensitivity of FAIRE-Seq Local changes in Chromatin nucleosome ATAC-Seq nucleosome accessibility occupied vs. MNAse-Seq positioning. depleted regions Measuring mobility Global decrease in Dynamics of of core histones and the mobility of chromatin binding chromatin binding FRAP nucleosomes and proteins. chromatin binding proteins. Changes in TADs Spatial genome Defining physical and compartments organization entities by contact Hi-C throughout the frequency. genome. Identifying electron- Ultrastructure of the lucent vs. electron- Emergence of nucleus dense regions in the Electron microscopy electron-dense nucleus regions.

A) Histone modifications, variants and chromatin associated proteins

During development, histone modifications are dynamically regulated at individual loci, as well as on a global scale. An example for local changes is H3K27me3 which decorates developmental genes. Prior to differentiation, Polycomb Repressive Complex

2 (PRC2) deposits H3K27me3 and represses lineage genes (Cao et al. 2002; Prezioso

11 and Orlando 2011; Shan et al. 2017). The presence of H3K4me3, an activating mark, on genes occupied by H3K27me3 in embryonic stem cells (ESCs) has been controversial. It has been proposed that these bivalent domains keep lineage genes in a poised state capable of rapid activation upon differentiation cues (Bernstein et al. 2006; Pan et al.

2007; Lien et al. 2011; Vastenhouw and Schier 2012). However, bivalent domains may only be present in ESCs, which are kept in an artificial state of permanent pluripotency. It is thought that they may exist only transiently or to a lesser extent in pluripotent epiblasts in embryos (Voigt, Tee, and Reinberg 2013).

Histone modifications are also regulated globally during development. For instance in zebrafish, frogs and flies, many histone modifications are low in early embryos and increase with the onset of zygotic transcription at the mid-blastula transition (Akkers et al.

2009; Lindeman et al. 2011; Schneider et al. 2011; Vastenhouw and Schier 2012; Li et al. 2014; K. Yuan and O’Farrell 2016).

For other systems, a global increase is observed only for specific modifications. In mouse embryos, the amount of H3K9me increases during differentiation. In particular,

H3K9 di-methylation levels increase significantly between the 2- and 4-cell stages (Yeo et al. 2005; Lepikhov and Walter 2004). H3K9 tri-methylation is present at the earliest stages of embryogenesis, but global levels also increase from the 16 cell to the blastocyst stage (Puschendorf et al. 2008).

It is possible that regulation of H3K9 methylation during development drives cell fate restriction, since H3K9 methylation can act as an epigenetic barrier against reprogramming cells into a pluripotent state (Becker, Nicetto, and Zaret 2015).

Alternatively, low amounts of H3K9 methylation in early embryos may provide a brief

12 window where repetitive sequences are transcribed, which is a pre-requisite for initiating silencing (Moazed 2009). In support of this notion, mutation of H3 Lysine 9 into Arginine leads to increased expression from piRNA clusters and transposons (Penke et al. 2016) and H3K9 methylation is enriched on repetitive elements at later stages of embryogenesis

(Zeller et al. 2016; McMurchy et al. 2017).

B) Chromatin accessibility

Nucleosomes are repositioned during differentiation of ESCs (Teif et al. 2012; W.

Zhang et al. 2016). In particular, nucleosome occupancy changes over key regulatory regions, including enhancers and binding sites of pluripotency factors (West et al. 2014), leading to a shift in transcriptional programs that drive differentiation. For instance, enhancers associated with pluripotency factors have on average lower nucleosome occupancy in ESCs compared to differentiated cell types (West et al. 2014). Similarly, in model organisms such as flies and worms, changes in nucleosome occupancy reflect initiation of specific developmental programs (Thomas et al. 2011; Daugherty et al. 2017).

In fly embryos, most of the changes in chromatin accessibility occur during the transition from stage 11 to 14, a period that coincides with extensive differentiation and cell-type specific gene activation (Thomas et al. 2011). As worm embryos hatch and develop into adulthood, accessibility at genes involved in embryonic morphogenesis and cell fate specification decrease, whereas the accessibility of genes involved in locomotion and larval development increase (Daugherty et al. 2017). All studies point towards local changes in nucleosome positioning to regulate gene expression.

It remains to be seen whether chromatin accessibility is altered globally during development, like histone modifications are. In genome-wide nucleosome mapping

13 assays, it is common practice to normalize different samples to the same sequencing coverage. This normalization works well for qualitative comparison, such as making peak calls, but may miss quantitative differences in nucleosome stability.

C) Dynamics of chromatin binding

High mobility of core histones in early mouse embryos or embryonic stem cells is thought to enable plasticity of gene expression and is gradually lost as silent heterochromatin domains form during differentiation (Meshorer et al. 2006; Bošković et al. 2014). FRAP assays show that mouse embryos lose histone mobility between the 2- cell and 8 cell stages (Bošković et al. 2014) and differentiating ESCs gradually lose H2B and H3 mobility (Meshorer et al. 2006). Similarly, mobility of linker histone H1 is restricted during development and this process requires histone modifications such as H3K9me

(Melcer et al. 2012). Maintenance of stable heterochromatin domains also involves transient binding and exchange of HP1 from chromatin, and the residence time correlates with the level of compaction (Cheutin et al. 2003; Meshorer et al. 2006).

While FRAP is good at tracking global changes, a disadvantage is that it merges two populations of histones: those bound to DNA and those free in the nucleoplasm.

Therefore, the dynamic nucleosomes observed by FRAP could reflect a larger pool of unbound histones in undifferentiated cells compared to cells of later stages.

D) Spatial genome organization

During ESC differentiation, TADs are mostly conserved, whereas active and inactive compartments are altered throughout the genome (Dixon et al. 2015). The inactive compartment expands in differentiated MS cells or IMR90 fibroblasts (Dixon et

14 al. 2015), cell types that typically gain repressive histone modifications as they differentiate (Hawkins et al. 2010; Xie et al. 2013).

Establishment of enhancer-promoter interactions through chromatin looping is crucial for the regulation of Homeotic gene (Hox) clusters that orchestrate anterior- posterior body axis and patterning during development (Andrey et al. 2013). During development, enhancer-promoter interactions are dynamically regulated in a cell-type specific manner concomitant to gene expression (Bonev et al. 2017; Rubin et al. 2017;

Freire-Pritchett et al. 2017).

In fly embryos, spatial organization of the genome is established during a small window that coincides with activation of zygotic transcription (Hug et al. 2017).

Interestingly, TADs are established independently of transcription, but depend on the transcription factor Zelda for insulation of boundaries (Hug et al. 2017).

E) Ultrastructure of the nucleus

In electron microscopy, electron dense regions (EDRs) in the nucleus are interpreted as heterochromatin. In animal cells, EDRs are rare during the earliest stages of embryogenesis, when cells are undifferentiated and transcriptionally quiescent, and become apparent as cells differentiate (Fadloun, Eid, and Torres-Padilla 2013; Politz,

Scalzo, and Groudine 2013). For example, Drosophila pole cells lack EDRs in early, pre- blastoderm stages but gain these structures upon differentiation (Mahowald 1968). In early mouse embryos, pluripotent epiblast cells have dispersed chromatin fibers, whereas differentiating trophectoderm and primitive endoderm possess regions of compacted chromatin (Ahmed et al. 2010; Park et al. 2004; Lessard and Crabtree 2010). Embryonic

15 stem (ES) cells recapitulate these embryonic transitions with the generation of heterochromatic puncta upon differentiation (Niwa 2007; Meshorer 2008).

• Facultative vs. Constitutive Heterochromatin

While heterochromatin is associated with certain marks and proteins, its definition remains ambiguous. Traditionally, heterochromatin is divided into two subtypes: facultative and constitutive. Facultative heterochromatin is present at gene-rich regions that can switch states between euchromatin and heterochromatin. A typical example for facultative heterochromatin is Polycomb Repressive Complex 2 (PRC2) depositing

H3K27me3 to repress lineage genes prior to differentiation (Cao et al. 2002; Prezioso and Orlando 2011; Shan et al. 2017). Constitutive heterochromatin, on the other hand, assembles mainly on centromeric, pericentromeric and telomeric regions known to harbor repetitive sequences enriched in H3K9 and DNA methylation (Saksouk, Simboeck, and

Déjardin 2015). However, recent findings suggest facultative and constitutive heterochromatin are not sufficient to describe the complexity of chromatin organization.

There is significant overlap in the distribution of H3K27me3 and H3K9me3 (Liu et al.

2011), and constitutive heterochromatin is not static during development. In differentiated cells, constitutive heterochromatin marked by H3K9me3 is densely packed, whereas in reprogrammed pluripotent cells, these regions are re-organized into dispersed 10nm chromatin fibers (Fussner et al. 2011). In sum, traditional definitions in chromatin organization may not be as black and white as once thought.

• Function of chromatin re-organization during development

Examples discussed above have led to the view that dispersed chromatin is a feature of pluripotent cells, and that the formation of heterochromatin domains characterize the differentiated state. It has been proposed that chromatin accessibility is critical for the pluripotent state in early embryos, where a broad spectrum of transcriptional programs need to be available. During differentiation, generation of heterochromatin is thought to focus transcriptional machinery on lineage genes and restrict cell fate potential (Gaspar-Maia et al. 2011). This notion is supported by the fact that certain factors involved in chromatin regulation contribute to pluripotency and differentiation, such as the Nucleosome Remodeling and Deacetylase Complex (NuRD),

Polycomb and Chd1 (Gaspar-Maia et al. 2009; Yuzyuk et al. 2009; Tursun et al. 2011;

Margueron and Reinberg 2011; Hu and Wade 2012; Custer et al. 2014; Zuryn et al. 2014;

T. Chen and Dent 2014).

Emergence of compact chromatin during differentiation influences replication timing as well. Early and late replicating domains show features of euchromatin and heterochromatin, respectively (Hiratani et al. 2010). Cell cycle is relatively fast at the earliest stages of embryogenesis and slows down during differentiation, coinciding with the emergence of heterochromatin (Boward, Wu, and Dalton 2016). Lack of heterochromatin in early embryos may facilitate rapid replication, or conversely, rapid replication may prevent heterochromatin formation.

• Non-genetic roles for heterochromatin

An emerging view is that heterochromatin has non-genetic functions beyond gene regulation and cell fate, and is important for:

i) Genome stability: Heterochromatin is thought to protect the genome from DNA damage, and regulated de-condensation is important for repair (Cann and Dellaire 2011;

Feng et al. 2016). In support of this notion, early mouse embryos that lack heterochromatin are extremely sensitive to X-rays (Goldstein, Spindle, and Pedersen

1975). Moreover, it has been proposed that HP1 accumulates at sites of DNA damage to protect cells from further damage (Luijsterburg et al. 2009; Gursoy-Yuzugullu, House, and

Price 2016). Recent work has demonstrated that heterochromatin also prevents transcription of repetitive sequences, which is a source of genome instability (Kim and

Jinks-Robertson 2012; McMurchy et al. 2017).

ii) Structural Robustness: It has been proposed that heterochromatin provides structural robustness to the nucleus to withstand mechanical forces in the cell. For instance, loss of heterochromatin diminishes the ability of the nucleus to withstand forces of the contracting heart (Furusawa et al. 2015). Another example for mechanical stress is cell migration, where cells pass through narrow constrictions (Gabi Gerlitz and Bustin

2011). Migrating melanoma cells have increased H3K9me3 and H3K27me3, marks associated with heterochromatin, and chromatin decondensation inhibits rate of cell migration in a transcription independent manner (G. Gerlitz and Bustin 2010).

iii) Night vision: There is a correlation between heterochromatin organization and night vision. In most cells, heterochromatin localizes to the nuclear periphery (Bank and

Gruenbaum 2011). In rod photoreceptor cells, heterochromatin is at the center of the nucleus (Solovei et al. 2009). It is thought that this inverted pattern results in a higher refractive index at the center of the nucleus, reduces light scattering and focuses the light on the photoreceptor plane, suggesting a physical role for genome organization.

1.3 SETDB1 mediated H3K9 methylation

• H3K9 methylation states

H3K9 methylation is broadly classified as a heterochromatin mark. H3K9 can be methylated to carry a single, double, or triple methyl group (me1, me2, me3). The degree of methylation can affect function; H3K9me2 and H3K9me3 are thought to have distinct roles. For instance, H3K9me2 is associated with gene silencing in euchromatin, whereas

H3K9me3 localizes to pericentric heterochromatin (Rice et al. 2003). Moreover, a recent study in the fission yeast S. pombe has revealed that transcriptionally active H3K9me2 domains precede RNAi-mediated silencing to establish silent H3K9me3 domains (Jih et al. 2017).

S. pombe has a single enzyme (Clr4) that generates all three forms, and is responsible for heterochromatin nucleation, spreading and maintenance (K. Zhang et al.

2008). In vertebrates, H3K9 methylation plays more diverse roles and relies on several partially redundant H3K9 methyltransferases that can direct different degrees of methylation (Allis and Jenuwein 2016):Suv39h1/2, G9a, GLP, SETDB1/2, Prdm3/16

(Kouzarides 2007; Pinheiro et al. 2012). Prdm3/16 exhibit mono-methyltransferase activity (Pinheiro et al. 2012), and H3K9me1 is converted into H3K9me3 by Suv39h1/2

(Peters et al. 2001; Rice et al. 2003). G9a is responsible for mono and di-methylation in

19 euchromatic regions (Rice et al. 2003; Tachibana et al. 2002), is embryonic lethal

(Tachibana et al. 2001) and de-regulated in cancer (Casciello et al. 2015). SETDB1 catalyzes mono- and di-methylation on H3K9 alone, and it is controversial whether it can also catalyze tri-methylation in the presence of binding partners (Wang et al. 2003;

Basavapathruni et al. 2016). There is functional cooperation between SETDB1, G9a and

Suv39h1, which participate in a multimeric protein complex (Fritsch et al. 2010).

• Developmental regulation of SETDB1

Chromatin organization by SETDB1 is crucial for development: Loss of SETDB1 results in the earliest death during embryogenesis amongst other H3K9 methyltransferases, at peri-implantation (Dodge et al. 2004). At the earliest stages of development when cells are uncommitted and pluripotent, SETDB1 represses developmental genes (Bilodeau et al. 2009) and maintains self-renewal in embryonic stem cells (ESCs) (Cho et al. 2012). At later stages of development, SETDB1 is required for trophoblast, hematopoetic, myogenic, osteoblastic differentiation and lineage restriction (P. Yuan et al. 2009; Koide et al. 2016; Song, Choi, and Lee 2015; Lawson et al. 2013). In fact, SETDB1 mediated H3K9 methylation acts as an epigenetic barrier for reprogramming differentiated cells into induced pluripotent stem cells (iPSCs) (J. Chen et al. 2013), suggesting it plays a critical in chromatin re-organization during development.

SETDB1 is maternally deposited and has a diffuse nuclear staining pattern at the

2 -cell stage. Once maternal stocks are depleted around the morula stage, SETDB1 reappears as punctate signals at the blastocyst stage (Cho, Park, and Kang 2011).

Interestingly, SETDB1 localizes to both the cytosol and the nucleus and shows activity in

20 both compartments (Loyola et al. 2006; Cho, Park, and Kang 2013). SETDB1 methylates histones in the cytosol during translation (Rivera et al. 2015) and in the nucleus in complex with heterochromatin factors like HP1 (Loyola et al. 2009).

Precise regulation of SETDB1 activity is important for not only development, but also overall organismal health: SETDB1 is overexpressed in Huntington’s Disease (Ryu et al. 2006), schizophrenia (Chase et al. 2013), and many cancer types such as lung cancer (Rodriguez-Paredes et al. 2014), gliomas (Spyropoulou et al. 2014) and melanomas (Ceol et al. 2011). Mis-regulation of SETDB1 is associated with an increase in the invasive potential of tumors in liver (Wong et al. 2016), prostate (Sun et al. 2014) and breast cancer (H. Zhang et al. 2014). Moreover, SETDB1 overexpression promotes survival of cancer cells after chemotherapy (Guler et al. 2017). One study postulates that

SETDB1 contributes to the oncogenic state by suppressing transposable elements in cancer cells, which helps immune evasion (Cuellar et al. 2017).

1.4 C. elegans embryogenesis as a model for chromatin organization

Many animals re-build heterochromatin domains during embryogenesis, including mice, flies and worms (Politz, Scalzo, and Groudine 2013; Yuzyuk et al. 2009). C. elegans embryos develop within 14 hours, with initiation of zygotic transcription at the 4-cell stage and gastrulation commencing at the 28-cell stage (Schauer and Wood 1990; Seydoux and Fire 1994; Sulston et al. 1983). Prior to gastrulation, cells are developmentally plastic, and their normal pattern of development can be reprogrammed by expression of selector genes ectopically, but this flexibility is lost during gastrulation (Mango 2009; Horner et al.

1998; Djabrayan et al. 2012; Zhu et al. 1998; Gilleard and McGhee 2001; Fukushige and

Krause 2005; Priess and Thomson 1987; Wood 1991; Sulston et al. 1983). Their rapid development, transparency and ex utero survival even at the earliest developmental stages, combined with the genetic tools available, makes C. elegans embryos an ideal system to study chromatin organization during differentiation.

The morphology of worm chromatin throughout embryogenesis has been tracked using artificial chromosomes that can be maintained indefinitely as chromatinized arrays of DNA (P Meister, Mango, and Gasser 2011; Yuzyuk et al. 2009; Fakhouri et al. 2010).

Prior to gastrulation, artificial chromosomes have a distended chromatin configuration and active RNA polymerase II (Yuzyuk et al. 2009). As gastrulation commences, artificial chromosomes compact, lose active RNA polymerase and associate with the nuclear lamina (Peter Meister et al. 2010; Fakhouri et al. 2010; Yuzyuk et al. 2009; P Meister,

Mango, and Gasser 2011). These behaviors suggest a dynamic nuclear environment during the transition from plasticity to cell fate acquisition.

Inactivation of several canonical heterochromatin proteins has no known effect on the pre-gastrula C. elegans embryo (Table 1.3; (Coustham et al. 2006; Paulsen,

Capowski, and Strome 1995; Capowski et al. 1991; Andersen and Horvitz 2007; Yuzyuk et al. 2009)). In addition, early embryos are particularly sensitive to DNA damaging agents such as UV irradiation. These observations suggest that early worm embryos bear reduced or altered heterochromatin (Hartman 1984; O’Neil and Rose 2006).

C. elegans embryos inherit many histone modifications including H3K27me3

(Arico et al. 2011; Bean, Schaner, and Kelly 2004; Gaydos, Wang, and Strome 2014;

Samson et al. 2014; Rechtsteiner et al. 2010), but H3K9me is erased in the mother’s

22 germ line during meiosis (Bessler, Andersen, and Villeneuve 2010), suggesting that the heterochromatin mark could be re-established during embryogenesis.

Table 1.3. List of chromatin factors without effects on the pre-gastrula embryo in C. elegans. Worm Gene Function Ortholog met-2 H3K9me1/me2 SETDB1 methyltransferase set-25 H3K9me3 G9a, SUV39 methyltransferase mes-2 H3K27me3 PCR2 E(z) methyltransferase cec-4 H3K9me binding protein -chromo hpl-2 H3K9me binding protein HP1 lin-61 H3K9me binding protein -L MBT2 lem-2 Nuclear Lamina protein MAN1 emr-1 Nuclear Lamina protein Emerin

In worms, MET-2 is the only enzyme that deposits H3K9me1/me2 and is orthologous to SETDB1 (Poulin et al. 2005; Andersen and Horvitz 2007). SET-25 mediated H3K9me3 is partially dependent on H3K9me2, and thus MET-2 is required for half of H3K9me3 (Towbin et al. 2012). Chromatin organization by MET-2 has roles in diverse biological processes such as repeat silencing, transgenerational inheritance of small RNAs (Lev et al. 2017), germline immortality (Andersen and Horvitz 2007), and mitochondrial stress (Tian et al. 2016). Unlike higher vertebrates, loss of MET-2 or SET-

25 does not result in embryonic lethality and makes worms an ideal system to study

H3K9me and its role in heterochromatin formation.

It is possible that met-2 and set-25 mutants are viable because loss of H3K9me is compensated by deposition of H3K27me by PRC2. A similar mechanism may exist in mammals, albeit not to the same extent, where loss of H3K9 has severe consequences.

Knockout of H3K9 methyltransferase Suv39h in mouse cells leads to increased PRC2

23 recruitment (Cooper et al. 2014), suggesting there is compensation under special circumstances.

In other systems, H3K9me leads to DNA methylation (Shankar et al. 2013;

Fadloun, Eid, and Torres-Padilla 2013; Andersen and Horvitz 2007). Worms lack cytosine methylation, however, indicating that H3K9me has roles beyond modifying DNA. In C. elegans, H3K9me is involved in transcriptional repression (Zheng et al. 2013; Kerr et al.

2014; Andersen and Horvitz 2007) and bound by the malignant-brain-tumor-repeat proteins LIN-61, MBTR-1 and the HP1-like protein HPL-2 (Koester-Eiserfunke and

Fischle 2011; Wirth et al. 2009; Garrigues et al. 2014). It is enriched on transcriptionally quiescent regions such as the X chromosome or regions associated with the nuclear lamina protein LEM-2 (Ikegami et al. 2010; Liu et al. 2011; Gerstein et al. 2010; Bean,

Schaner, and Kelly 2004; Bessler, Andersen, and Villeneuve 2010). Its enrichment on repetitive sequences such as transposons prevents their transcription and is crucial for genome integrity (Zeller et al. 2016; McMurchy et al. 2017).

The motivation of this study is to better understand the molecular underpinnings of chromatin re-organization during embryogenesis. In this thesis, I focused on the regulation of the H3K9 methyltransferase MET-2, which is required for generating heterochromatin in C. elegans embryos. First, I identified two conserved regulators of

MET-2 that are required for H3K9me accumulation during development and investigated their role in H3K9 methylation (Chapter 2). Second, I studied the determinants that influence timely acquisition of the mark in developing embryos (Chapter 3). My studies reveal that nuclear accumulation of MET-2 and its regulators dictates the timing of

H3K9me and heterochromatin domains during embryogenesis.

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CHAPTER 2

REGULATION OF A HISTONE METHYLTRANSFERASE TIMES THE ONSET OF

HETEROCHROMATIN FORMATION IN C. ELEGANS EMBRYOS

This chapter is currently under review for publication.

Beste Mutlu, Huei-Mei Chen, James J. Moresco, Barbara D. Orelo, Bing Yang, John M.

Gaspar, Sabine Keppler-Ross, John R. Yates III, David H. Hall, Eleanor M. Maine, Susan

E. Mango.

2.1 Abstract

Heterochromatin formation during early embryogenesis is timed precisely, but it has been elusive how this process is regulated. Here we report the discovery of a histone methyltransferase complex whose nuclear accumulation determines the onset of heterochromatin formation in C. elegans embryos. We find that the inception of heterochromatin generation coincides with the accumulation of the methyltransferase

MET-2 (SETDB) into nuclear hubs. The absence of MET-2 results in delayed and disturbed heterochromatin formation, whereas accelerated nuclear localization of the methyltransferase leads to precocious heterochromatin. We identify two factors that bind to and function with MET-2: LIN-65, which resembles ATF7IP, localizes MET-2 into nuclear hubs, and ARLE-14, orthologous to ARL14EP, promotes stable association of

MET-2 with chromatin. These data reveal that nuclear accumulation of MET-2 in conjunction with LIN-65 and ARLE-14 regulates timing of heterochromatin domains during embryogenesis.

2.2 Main text

The nucleus of a young embryo undergoes major reorganization as it transitions from fertilized egg to multicellular embryo. As cells acquire specific fates and zygotic transcription commences, the nucleus is segregated into distinct domains of euchromatin and heterochromatin (Politz, Scalzo, and Groudine 2013). While much has been learned about the mechanisms that control cell-fate specification and the onset of zygotic transcription (Guven-Ozkan et al. 2008; Lee, Bonneau, and Giraldez 2014), little is understood about the processes that establish chromatin domains de novo during embryogenesis. To begin to tackle this question, we examined heterochromatin formation in the nematode C. elegans.

We began our analysis with a survey of wild-type embryos using two assays for heterochromatin: First, we used transmission electron microscopy (TEM), where heterochromatin domains can be detected as electron-dense regions (EDRs) within nuclei (Davies 1968; Hall, Hartwieg, and Nguyen 2012). Second, we surveyed histone modifications to track their abundance and morphology in early embryos. By TEM, nuclei at the earliest stages appeared relatively homogenous, with light speckling in the nucleoplasm and a nuclear envelope free of electron-dense material (Figure 2.1A-B).

Upon initiation of gastrulation (~21-50 cell stage), embryos gained more electron-dense puncta throughout their nuclei. By mid-gastrulation (51-100 cell stage), dark material was observed abutting the nuclear envelope, and the nucleoplasmic puncta coalesced into larger, but fewer, electron-dense compartments. These features became more pronounced over time, with large EDRs that spanned the nucleus and bordered the nuclear periphery (Figure 2.1B, >200 cells). We note that EDRs appeared throughout the

51 embryo, suggesting that cells destined to produce different cell types nevertheless generated heterochromatin at about the same time in development (Supplementary

Figure 2.1A).

Antibody staining revealed a dramatic increase in Histone H3 Lysine 9 methylation

(H3K9me) from fertilization to the mid-gastrula in interphase cells. H3K9me2 increased ten-fold (p=3.92x10-19), while H3K9me1 (p=0.001) and H3K9me3 (p=0.001) each increased two-fold (Figure 2.1C, D). H3K9me2 was barely detectable at fertilization, but bright puncta became apparent by the 20-cell stage throughout nuclei. The signal intensified during gastrulation, with more puncta and brighter staining within puncta (~51-

100-cell stage, mid-stage). A time series of whole embryo stains indicated that most interphase nuclei behaved similarly (Supplementary Figure 2.1B), and two different antibodies against H3K9me2 gave identical results (Supplementary Figure 2.1C). Both puncta of histone modifications and EDRs arise during gastrulation, suggesting H3K9me stains are a useful proxy to visualize heterochromatin domains. We note that not all histone modifications changed in early embryos: neither levels nor morphology of

H3K27me3 or pan-H4Ac were altered (Figure 2.1C, D; see also (Bender et al. 2004)).

To identify the molecular basis of heterochromatin and H3K9me formation, we focused on the methyltransferase MET-2. MET-2 is homologous to vertebrate SETDB1

(Poulin et al. 2005; Andersen and Horvitz 2007), and required for virtually all H3K9me1 and H3K9me2 (Bessler, Andersen, and Villeneuve 2010; Towbin et al. 2012). H3K9me2 was regulated dynamically during embryogenesis, similar to EDRs and its location in the genome, by ChIP, tracks well with heterochromatin proteins such as HPL-2/HP1

Figure 2.1. Heterochromatin and H3K9me domains are established during embryogenesis. A. Time line of C. elegans embryogenesis. Stages are color-coded: light green (<20 cell, pre-gastrula), green (21-200 cell, gastrula), dark green (200-500 cell, late stage). Morphogenesis starts after the 500-cell stage and was not analyzed in this study. B. Transmission electron micrographs (TEM) of representative nuclei from wild-type embryos (Scale bar, 1 µm). C. Survey of histone modifications. Representative single nuclei at designated embryonic stages stained for histones and DNA (Scale bar, 2 μm). D. Quantitation of histone modifications normalized to total histone H3.

(Garrigues et al. 2014). Thus, we asked whether loss of MET-2 impacted heterochromatin domains visible by TEM. To control for TEM fixation and sectioning, we examined cytoplasmic organelles and yolk droplets from met-2 mutants, which resembled those of wild-type embryos (Supplementary Figure 2.2A, B). Pre-gastrula met-2 embryos matched their wild-type counterparts, with homogeneous, translucent nuclei (Figure 2.2A).

However, the speckles observed in wild-type nuclei at the 20-cell stage were dimmer in met-2 mutants, and they failed to coalesce into EDRs by the 50-100-cell stage (Figure

2.2A). EDRs emerged in older embryos, but they occupied less nuclear volume and were reproducibly paler (Figure 2.2A). We performed line-scan analysis to quantify the appearance of nuclear EDRs. The standard deviation of line-scan values is higher in nuclei with EDRs compared to homogeneous nuclei because of the dark EDRs in pale nucleoplasm. met-2 nuclei had a homogeneous distribution of signal and a smaller standard deviation at every stage (Figure 2.2B). These results indicate that met-2 is critical for the timely formation of segregated heterochromatin domains.

Given the dependence of H3K9me2 on MET-2, we asked if expression of MET-2 tracked with the onset of H3K9me2. We found that MET-2 protein gradually shifted from the cytosol to the nucleus, from the 2-cell stage to the onset of gastrulation, and this change was observed for both endogenous MET-2 and single-copy MET-2 reporters

(Figure 2.2C-E, Supplementary Figure 2.2D). In 1-4-cell embryos, MET-2 was distributed throughout nuclei and cytoplasm with little nuclear accumulation. As embryos aged, the concentration of MET-2 within nuclei increased approximately five-fold (Figure 2.2D, p=0.0002). The absolute level of MET-2 protein did not change significantly over time

(Figure 2.2E). We note that early embryos acquired nuclear MET-2 and H3K9me2

54 transiently during prophase (Supplementary Figure 2.2E, F), but we focus on interphase nuclei here.

Given that met-2 is necessary for heterochromatin domains, and H3K9me2 accumulated dynamically, we asked whether regulation of MET-2 constituted part of the embryonic timer for heterochromatin establishment. We hypothesized that if nuclear

MET-2 was rate-limiting, then premature accumulation of MET-2 in nuclei would lead to precocious H3K9me2 and initiate heterochromatin. We added a nuclear localization signal (NLS) from c-Myc to a FLAG-tagged copy of endogenous MET-2 using Crispr, which increased nuclear MET-2 by approximately two-fold in pre-gastrula embryos

(Figure 2.2F, I). Increased nuclear MET-2 led to precocious accumulation of H3K9me2, beginning at least one cell division earlier than wild-type embryos (Figure 2.2G, H, J; p<0.05 at all stages). These results suggest that gradual accumulation of MET-2 within nuclei initiates H3K9me2.

To understand how MET-2 is regulated, we searched for binding partners using immunoprecipitation followed by Multidimensional Protein Identification Technology Mass

Spectrometry, using MET-2::GFP and 3xFLAG::MET-2. Wild-type C. elegans bearing no tagged proteins, and strains bearing an unrelated GFP or FLAG reporter served as negative controls. We chose a candidate list of interacting partners based on the specificity of MET-2 binding, on the peptide counts and protein coverage. Seventeen candidates were chosen for secondary screening for their effects on H3K9me. From this survey, two proteins emerged as likely MET-2 partners (Supplementary Figure 2.3A): LIN-

65 is a 100kDa protein and the most abundant interactor of MET-2 (Supplementary Figure

2.3B); B0336.5 codes for a smaller protein (~30kDa) and had lower spectral counts, but

Figure 2.2. Nuclear accumulation of H3K9 methyltransferase MET-2/SETDB1 determines the onset of heterochromatin formation. A. TEM of representative nuclei from wild-type (WT) or met-2 embryos (Scale bar, 1 µm). B. TEM line-scan analysis for WT (black) or met-2 (magenta) nuclei and the standard deviation (stdev). Line scan for a single nucleus is shown as an example.

Figure 2.2 continued C. Embryos stained for MET-2::GFP (upper) and HIS-72::mCherry (lower) at designated stages of embryogenesis (Scale bar, 2 μm). D-E. Quantitation of nuclear (D) and total (E) MET-2 at designated stages, normalized to HIS-72::mCherry. F-G. Localization of 3xFLAG::MET-2 with a c-Myc NLS compared to 3xFLAG::MET-2 control (F) and corresponding H3K9me2 levels (G). H. Interphase nuclei showing H3K9me2 levels for the c-myc NLS construct compared to 3xflag::met-2 embryos. I. 3xFLAG::MET-2 line scans in pre-gastrula embryos (1-4 cell stage) with (red) or without (grey) the c-myc NLS. Average of line scans across multiple nuclei are shown and error bars denote standard error of the mean. J. H3K9me2 levels normalized to H3 for the NLS construct (red) compared to control embryos (grey).

similar protein coverage as LIN-65 (Supplementary Figure 2.3A). We renamed B0336.5 as arle-14 for ARL14 Effector Protein, as explained below.

To test the role of MET-2 binding partners in H3K9me deposition, we analyzed loss-of-function mutants. lin-65 mutants had reduced levels of H3K9me1 and H3K9me2, and low, dispersed H3K9me3, similar to met-2 mutants (Figure 2.3A, B). arle-14 mutants resembled a partial loss of met-2 activity, with reduced H3K9me1/me2 levels and largely normal H3K9me3 (Figure 2.3A, B).

lin-65 and arle-14 resembled met-2 mutants in two additional assays. First, an important role of MET-2 is to silence repetitive DNA (Zeller et al. 2016; McMurchy et al.

2017). RNAs for two repeats were de-repressed by arle-14 mutations and, to a greater degree, by lin-65 (Figure 2.3C). Second, de-repressed repeats lead to a mortal germline phenotype for met-2 mutants at 26°C (Zeller et al. 2016). Similarly, lin-65 mutants became sterile after a single generation, and arle-14 mutants after two generations. These results indicate that LIN-65 and ARLE-14 are bona fide binding partners for MET-2 and contribute to its functions.

To address whether LIN-65 and ARLE-14 contribute to the onset of heterochromatin formation, we examined their expression during embryogenesis. Both proteins behaved similarly to MET-2: they were enriched in the cytoplasm from the one- cell stage through the eight-cell stage, but gradually moved into nuclei thereafter; total levels did not change (Supplementary Figure 2.3C, D). During gastrulation, we observed concentrated hubs of MET-2, LIN-65 and ARLE-14 emerge within nuclei (Figure 2.3D, E,

F). H3K9me2, MET-2, LIN-65 and ARLE-14 co-localized in many of these hubs and excluded the activating mark H3K4me3 (Figure 2.3G).

The antibody stains suggested that MET-2 could bind LIN-65 and ARLE-14 in either the cytoplasm or the nucleus, but imaging of proteins under the light microscope lacks the resolution to define where binding occurs. We took advantage of the Proximity

Ligation Assay (PLA), which detects pairs of proteins when they are within ~30 nm of each other, to investigate MET-2 binding to its partners. Positive and negative controls demonstrated that our PLA signals were specific (Supplementary Figure 2.3E). At the earliest stages of embryogenesis, we observed MET-2 PLA+ signal with LIN-65 and

ARLE-14 in the cytoplasm but rarely in the nucleus (Figure 2.3H, I, J). As embryos matured, we continued to detect PLA signals in the cytoplasm but also observed signal within nuclei for MET-2 with both LIN-65 and ARLE-14. These results reveal that MET-2 interacts closely with LIN-65 and ARLE-14 and all three proteins accumulate in nuclear hubs over time.

To address how ARLE-14 contributes to the deposition of H3K9me2 by MET-2, we examined MET-2 in wild-type and arle-14 embryos. Neither the localization nor level of

MET-2 changed in arle-14 mutants (Supplementary Figure 2.4A). By ChIP-seq, the

Figure 2.3. MET-2 and two conserved binding partners form concentrated nuclear hubs at gastrulation. A. Whole embryos stained with antibodies against methylated H3K9 and pan-histones in wild-type vs. met-2, lin-65 or arle-14 mutants. Scale bar, 2 μm. B. H3K9me levels in mutant embryos normalized to on-slide control embryos and to pan- histone. C. Expression of CEREP4 and CEMUDR1 repeat RNAs by RT-qPCR in wild-type vs. met-2, lin-65 or arle-14 mutant embryos. D-F. Representative singe nuclei showing 3xFLAG::MET-2 (D), LIN-65::3xFLAG (E) and ARLE-14 (F) localization at different embryonic stages (Scale bar, 2 μm). Quantitation of signal intensity in nuclear hubs (red), nuclear regions excluding hubs (“non-hub”, dark grey) and cytosol (light grey). G. Single nuclei showing 3xFLAG::MET-2, LIN-65::3xFLAG and ARLE-14 co-localization in concentrated protein hubs and exclusion of activating mark H3K4me3 (Scale bar 2 μm). H. PLA signal showing interactions between MET-2/LIN-65 and MET-2/ARLE-14.

Figure 2.3 continued I-J. Percentage of nuclear PLA dots per total dots in embryos for MET-2/LIN-65 and MET- 2/ARLE-14 interactions.

distribution of H3K9me2 was also normal in arle-14 mutants (wild-type vs. arle-14 gave a genome-wide correlation of 0.8 and 0.92 for two independent experiments) (Figure 2.4A).

Thus, ARLE-14 is not required to target MET-2 in the genome (Figure 2.4B). Rather, arle-

14 affected the degree of association of MET-2 with chromatin. Quantitative analysis by

ChIP-qPCR revealed a two-fold decrease in MET-2::GFP binding to known genomic targets in arle-14 mutants (Figure 2.4C). Moreover, loss of arle-14 delayed the accumulation of H3K9me2 in early embryos (Figure 2.4D, E), revealing that ARLE-14 is also critical for proper timing of accumulation.

Next, we examined LIN-65. In lin-65 mutants, MET-2 remained cytoplasmic (Figure

2.4F, Supplementary Figure 2.4B). MET-2 levels did not decrease in lin-65 mutants

(Supplementary Figure 2.4C), indicating that LIN-65 affected the subcellular distribution of MET-2 and not its stability. Thus, LIN-65 is required for the timely accumulation of MET-

2 within nuclei. We note that LIN-65 has additional roles after MET-2 and LIN-65 become nuclear, as bypassing the requirement for LIN-65 for nuclear MET-2 with a c-Myc NLS does not rescue H3K9 methylation (Figure 2.4H-K).

The NLS experiment suggested that nuclear accumulation of MET-2 might be rate- limiting for H3K9me2 and heterochromatin formation. To test this idea further, we reduced the dose of MET-2 or LIN-65 by examining embryos from met-2/+ or lin-65/+ heterozygotes. Embryos from met-2/+ mothers behaved like wild-type, with normal levels and distribution of MET-2 and H3K9me2 (Supplementary Figure 2.4D, E), suggesting that

MET-2 is dosage compensated. A half dose of lin-65 lead to reduced accumulation of

MET-2::GFP within nuclei and more cytoplasmic MET-2 (Supplementary Figure 2.4F).

H3K9me2 accumulated more slowly, and pre-gastrula embryos had approximately half the level of age-matched controls (Figure 2.4L-N p=1.5 x10-14). These results indicate that

LIN-65 is rate-limiting for nuclear accumulation of MET-2 and H3K9me.

This study makes two contributions towards understanding heterochromatin formation during embryogenesis (Figure 2.4P). First, we have identified two binding partners, LIN-65 and ARLE-14, that are critical for MET-2 to localize to nuclei and associate with chromatin. MET-2, LIN-65 and ARLE-14 likely bind in a complex because

MET-2 requires LIN-65 for nuclear localization, whereas ARLE-14 requires MET-2 for both nuclear localization and cellular accumulation (Supplementary Figure 2.4L).

lin-65 belongs to the synMuv B subclass of regulators, which are involved in chromatin regulation and transcriptional repression (Fay and Yochem 2007). Although lin-65 had been annotated as a novel protein (Ceol et al. 2006), we found similarities between LIN-65 and the co-factor ATF7IP (Activating Transcription Factor 7-Interacting

Protein). LIN-65 has a high-probability coiled-coil region predicted by PCOILS (CC; fig.

S4G) and a high-confidence beta-sandwich in the C-terminus, like ATF7IP (S; fig. S4H). fig. S4I). Like LIN-65, ATF7IP binds and localizes SETDB1 to nuclei (Wang et al. 2003;

Ichimura et al. 2005; Koch et al. 2009). However, LIN-65 is not an obvious orthologue of

ATF7IP and may be an example of convergent evolution.

Prior to this study, ARLE-14 was an uncharacterized C. elegans protein with homology to ADP Ribosylation Factor 14 Effector Protein (Supplementary Figure 2.4J).

Figure 2.4

Figure 2.4 continued

Figure 2.4. LIN-65 and ARLE-14 promote MET-2 nuclear localization and chromatin association. A-B. H3K9me2 ChIP-Seq (logLR) track in wild-type vs. arle-14 mutant embryos (chromosome III and loci: rep-1, Y22D7AL.7, grl-16). C. MET-2::GFP ChIP-qPCR for wild-type (green) vs. arle-14 (yellow) mutant embryos. Inset shows H3K4me3 ChIP-qPCR as a control. D-E. Acquisition of H3K9me2 in wild-type vs. arle-14 mutants during embryogenesis, with an H3 co-stain (D) and quantitation normalized to H3 (E). F. Distribution of MET-2::GFP in wild-type vs. lin-65 mutants vs. no-GFP wild-type strain (Scale bar, 2 μm). Note that this H3 antibody detects mostly cytosolic histone H3 during mitosis. G. Line-scan analysis across embryonic nuclei shows mean MET-2::GFP intensity in wild- type (green) vs. lin-65 (pink) mutants vs. no-GFP control (“-“, grey). Average of line scans across multiple nuclei are shown and error bars denote standard error of the mean. H. 3xFLAG::MET-2 with a c-Myc NLS in a wild-type and lin-65 mutant background. I. H3K9me2/me3 levels in embryos with a single copy ZEN-4::GFP (“WT”, on-slide control) or NLS::3xFLAG::MET-2 in a lin-65 mutant background. J-K. Quantitation of H3K9me2/me3 levels in wild-type (grey) or lin-65; NLS::3xflag::met- 2 (blue) embryos.

Figure 2.4 continued L,N. H3K9me2/me3 levels in embryos with a single copy ZEN-4::GFP (“WT”, on-slide control) or progeny of lin-65(+/-) heterozygous mothers identified by HIS-72::mCherry at designated stages of embryogenesis. M,O. Quantitation of H3K9me2/me3 levels from wild-type (grey) or lin-65(-/+) (purple) offspring. P. In early embryos, MET-2 (grey), LIN-65 (red) and ARLE-14 (yellow) are enriched in the cytosol, and there is little H3K9me2 or heterochromatin (light green). As embryos mature, MET-2 and interactors gradually accumulate in nuclei, form concentrated nuclear hubs and deposit H3K9me2. MET-2-dependent H3K9 methylation is required to generate heterochromatin domains (compacted, dark green).

In vertebrates, the only published function for ARL14EP is in the cytoplasm (Paul et al.

2011). However, the Protein Atlas shows ARL14EP in the nuclei of many human tissues

(www.proteinatlas.org). We surveyed large-scale interaction databases (Rolland et al.

2014; Giot et al. 2003; Guruharsha et al. 2011) and uncovered an interaction between

ARL14EP and SETDB proteins in both humans and Drosophila (Supplementary Figure

2.4K). These data suggest that, in addition to its cytoplasmic role, ARLE-14 and its orthologues share a conserved function in nuclei with SETDB methyltransferases.

Interestingly, human ARL14EP and H3K9 methylation have been implicated in polycystic ovary syndrome, but were considered independent aspects of the disease (Hayes et al.

2015; Kokosar et al. 2016; Eini et al. 2017). Our results suggest there may be a link between ARL14EP and H3K9 methylation in PCOS.

Second, we find that the onset of heterochromatin formation depends on the gradual accumulation of MET-2 within nuclei. Similar to C. elegans, mammals and

Drosophila rebuild heterochromatin domains during embryogenesis (Politz, Scalzo, and

Groudine 2013; Yuan and O’Farrell 2016). More generally, lack of heterochromatin domains appears to be a feature of undifferentiated cells, including embryonic stem cells

64 and planarian neuroblasts, and differentiation involves re-establishing heterochromatin

(Politz, Scalzo, and Groudine 2013). Examination of previous studies suggest murine

SetDB1 is cytoplasmically enriched in early embryos, but its function has been difficult to address due to early lethality (Dodge et al. 2004; Cho et al. 2012). An intriguing idea is that nuclear localization of SETDB with ATF7IP and ARL14EP initiates heterochromatin formation in other animals as well.

2.3 Materials and Methods

Strains. Strains were maintained at 20oC according to (Brenner 1974), unless stated otherwise.

N2 (wild-type Bristol)

RB1789 met-2 (ok2307) III, provided by the C. elegans Gene Knockout Project at

OMRF.

MT13232 lin-65(n3441) I (Ceol et al. 2006).

SM2078 stIs10389 (pha-4::gfp::3xflag); pha-4 (q500) rol-9 (sc148) (Hsu et al. 2015).

SM2333 pxSi01 (zen-4::gfp, unc-119+) II; unc-119(ed3) III.

SM2529 arle-14/B0336.5(tm6845) III, provided by the Japanese National Biosource

Project.

JAC500 his-72(csb43[his-72::mCherry]) III, provided by John Calarco (Norris et al.

2015).

EL597 omIs 1 [Cb-unc-119 (+) met-2::gfp II].

SM2491 omIs 1 [Cb-unc-119 (+) met-2::gfp II]; met-2(ok2307) unc-119 (ed3) III.

SM2533 omIs1 [Cb-unc-119 (+) met-2::gfp II]; arle-14(tm6845) III.

SM2536 omIs1 [Cb-unc-119 (+) met-2::gfp II]; lin-65 (n3441) I.

SM2532 [Cb-unc-119 (+) met-2::gfp II]; his-72(csb43[his-72::mCherry]) III.

SM2575 lin-65::3xflag I. This study.

EL634 3xflag::met-2 III. This study.

SM2576 arle-14(tm6845) III, lin-65::3xflag I. This study.

SM2580 NLS::3xflag::met-2 III. This study.

Number of experiments and embryos surveyed.

For imaging experiments, first, many embryos were surveyed under the microscope through the eye piece and general trends noted. Then a random subset of embryos was imaged and analyzed more deeply, with quantitation. Details of analysis are described separately in the Image analysis section, and the analysis gave the same qualitative result as the trends observed in the initial survey. Below are the numbers for the quantitation.

Fig. 2.1C. >50 embryos were surveyed for each histone mark. At least 5 wild-type embryos from each developmental stage were imaged and analyzed in N=3 experiments.

Fig. 2.2C. >100 embryos were surveyed. A total of 37 embryos were imaged and analyzed in N=3 experiments.

Fig. 2.2F. >50 embryos were surveyed for each strain.14 wild-type and 14 NLS embryos were imaged and analyzed in N=3 experiments.

Fig. 2.2G. >30 more embryos for each strain were surveyed. 15 wild-type and 24 NLS embryos were imaged and analyzed in N=3 experiments.

Fig. 2.3A. We analyzed the following number of embryos in N=3 experiments:

H3K9me1: WT vs. met-2(ok2307) (5, 7), WT vs. lin-65(n3441) (8, 12), WT vs. arle-

14(tm6845) (6, 7). H3K9me2: WT vs. met-2(ok2307) (11, 20), WT vs. lin-65(n3441)

(10, 13), WT vs. arle-14(tm6845) (22, 21). H3K9me3: WT vs. met-2(ok2307) (14, 12),

WT vs. lin-65(n3441) (6,6), WT vs. arle-14(tm6845) (5,6).

Fig. 2.3C. Error bars denote standard error of the mean for N=3 experiments.

Fig. 2.3H. For MET-2/ARLE-14 PLA, a total of 20 embryos were analyzed in N=3 experiments. For MET-2/LIN-65 PLA, a total of 28 embryos were analyzed in N=3 experiments.

Fig. 2.4A. 2 biological replicates were processed in parallel.

Fig. 2.4C. Error bars denote standard error of the mean for N=3 experiments.

Fig. 2.4D. >40 embryos were surveyed for each strain.10 wild-type and 19 arle-

14(tm6845) mutant embryos were imaged and analyzed in N=3 experiments.

Fig. 2.4F. >100 lin-65 mutant embryos were surveyed. 14 wild-type and 31 lin-65 mutant embryos were imaged and analyzed in N=3 experiments.

Fig. 2.4L. 22 wild-type and 25 lin-65 +/- embryos were analyzed in N=3 experiments.

Sup. Fig. 2.3C. 41 embryos were analyzed in N=3 experiments.

Sup. Fig. 2.3D. 30 embryos were analyzed in N=3 experiments.

Sup. Fig. 2.4A. >30 mutant embryos were surveyed. 31 wild-type embryos and 21 arle-

14(tm6845) mutant embryos were imaged and analyzed in N=3 experiments,

Sup. Fig. 2.4C. Error bars denote standard error of the mean for N=2 experiments.

Sup. Fig. 2.L. >50 embryos were surveyed for each strain.11 wild-type vs. 11 lin-65 mutant embryos, 8 wild-type vs. 4 met-2(ok2307) mutant embryos were imaged and analyzed in N=3 experiments.

Generation of MET-2, LIN-65, ARLE-14, ZEN-4 and HIS-72 reagents

MET-2: To generate EL634, we inserted the 3xFLAG tag

(DYKDHDGDYKDHDIDYKDDDDK) at the endogenous met-2 locus using Crispr (Paix et al. 2014; Arribere et al. 2014). The construct places the tag at the amino terminus of MET-

2 and is inserted immediately after the start codon without any linker sequences. To generate EL597, we inserted a single copy of MET-2::GFP with its endogenous promoter and upstream gene R05D3.2 at the ttTi5605 locus on chromosome II by MosSCI

(Frøkjaer-Jensen et al. 2008; Zeiser et al. 2011). The GFP tag was placed at the carboxyl terminus of MET-2 and was inserted immediately before the stop codon without any linker sequences. Both the Crispr construct and the MosSCI construct could rescue H3K9me2 deposition, although the Crispr allele did so better than the MosSCI allele. The MET-2 antibody was generated against the first 17 amino acids of endogenous MET-2 and affinity purified.

To generate SM2580 (NLS::3xFLAG::MET-2), the c-Myc NLS sequence

(CCAGCCGCCAAGCGTGTCAAGCTCGAC) was added directly upstream of

3xFLAG::MET-2 without any linker sequence by Crispr (Paix, Folkmann, and Seydoux

2017). For the insertion, the 3xFlag sequence in EL634 was targeted by the following guide RNA: ATGGACTACAAAGACCATGA(CGG). The dpy-10 locus was used as a phenotypic marker. Because it segregated independently from the met-2 locus, non-roller non-dpy worms were isolated for further analysis by single worm PCR and genotyping.

The edit was confirmed by sequencing the 200bp region around the insertion. crRNA, tracrRNA, and Cas9 protein were ordered from the IDT Alt-R genome editing system. The

97 bp repair template was synthesized and PAGE purified by IDT.

LIN-65: We inserted a 3xFLAG at the endogenous lin-65 locus using Crispr (Paix,

Folkmann, and Seydoux 2017). The sequence of the guide RNA was

TCATTCGAGAGTGATGAAGG(TGG). The 3xFLAG tag is located at the C-terminus of

LIN-65 and is inserted directly before the stop codon without any linker sequences. The dpy-10 locus was used as a phenotypic marker. Because it segregated independently from the lin-65 locus, non-roller non-dpy worms were isolated for further analysis by single worm PCR and genotyping. The edit was confirmed by sequencing the 200bp region around the insertion. crRNA, tracrRNA, and Cas9 protein were ordered from the IDT Alt-

R genome editing system. The 136 bp repair template was synthesized and PAGE purified by IDT.

ARLE-14: An antibody against endogenous ARLE-14 was generated. Bacteria containing arle-14 cDNA in a pET-47b(+) (Novagen, #71461) plasmid backbone were grown at 30oC for 19 hours in LB + 50μg/ml Kanamycin. The culture was diluted 1:5 in LB + Kan and protein expression was induced with 0.25mM IPTG at 30oC for 3 hours. Bacteria were pelleted, and flash frozen at -80oC. The pellet was resuspended in 20ml Lysis Buffer

(50mM Tris pH 7.2, 300mM NaCl, 5mM B-ME, 10% glycerol) and digested with 200μl of lysozyme (50mg/ml, Thermo Fisher Scientific #90082) for 30 minutes on ice. Following lysozyme digestion and sonication on ice (4 cycles, 30sec ON, 1min OFF. Output Control

3, Duty Cycle 50%, Pulsed), the protein was purified from inclusion bodies as follows: The sample was centrifuged at 4000rpm at 4oC for 15min and the supernatant discarded. The

69 pellet was resuspended in 20ml Lysis Buffer with 1% Triton-X and 200μl Turbo DNAse

(Thermo Fisher Scientific AM2239) and incubated for 20 min at room temperature. The sample was sonicated and centrifuged again with the same settings as before, and the supernatant discarded. The pellet was rinsed once with Dilution Buffer (10mM Tris/Cl pH

7.5,150mM NaCl, 0.5mM EDTA) and resuspended in Denaturation Buffer (50mM Tris-

HCl pH 8, 300mM NaCl, 2mM B-ME, 5mM MgCl2, 6M urea) by gentle rocking on a shaker at room temperature for 1 hour. The solution was dialyzed against 50mM Tris-Cl pH 8,

150mM NaCl, 5mM MgCl2 and 1mg/ml protein was sent to Covance for injections into rabbits. Total IgG purification was performed by Covance after the final bleed. For in vivo imaging, the antibody solution was pre-cleared overnight with arle-14(tm6845) mutant embryos before use. Protocol described in the antibody staining section was followed to prepare arle-14(tm6845) embryos and stain them with the ARLE-14 antibody. The resulting pre-cleared antibody solution was transferred to a fresh tube, stored at 4oC for

<1week and used in staining experiments.

ZEN-4: ZEN-4::GFP was amplified from bsem1129 (Von Stetina et al. 2017) with zen-4_uni_5'_nested_2_attB1

(GGGGACAAGTTTGTACAAAAAAGCAGGCTGCAAAAAGTCGCATCTGGGAA; attB1 underlined) and unc-54_3'UTR_Hobert_nested_3'_attB2

(GGAAACAGTTATGTTTGGTATATTGGGACCCAGCTTTCTTGTACAAAGTGGTCCCC

; attB2 underlined) primers using Takara PrimeStar (Von Stetina et al. 2017). The resulting attB-flanked PCR product was recombined into pCFJ151 (Addgene) using

Gateway BP Clonase II (Invitrogen/Thermo Fisher) to create bsem1267. SM2333 was

70 generated by injecting bsem1267 along with pCFJ601, pMA122, pGH8, pCFJ90 and pCFJ104 (all available from Addgene) into SM2288 (ttTiS605 II; unc-119(ed3)III). The mosSCI protocol on wormbuilder.org was used to generate single integrants (Frøkjaer-

Jensen et al. 2008). SM2333 was used as an on-slide wild-type control in antibody stains.

HIS-72: The mCherry tag was inserted at the C-terminus of the endogenous his-72 locus by Crispr (Norris et al. 2015). Briefly, JAC499 was injected with Cre recombinase to remove the selection cassette and produce a functional HIS-72::mCherry protein. JAC500 his-72(csb43[his-72::mCherry]) III was used as a histone control in MET-2::GFP stains

(Fig. 2C) and to mark cross-progeny after mating (Fig. 4H, S4E). The mCherry tag did not interfere with H3K9me2 (fig. S2C).

Antibody staining. Antibody staining was performed as described previously (Kiefer et. al, 2007). The following antibodies were used for immunostaining with 5min 2% paraformaldehyde (PFA), 3 min methanol (for all except ARLE-14 and MET-2, which was fixed with 10min 2% PFA, 3min methanol). For mutant configurations, an on-slide wild- type sample was included, marked with a single-copy ZEN-4::GFP tag. On-slide controls allowed better quantitation between different genotypes or stages.

H3K9me1 (1:200) Abcam ab8896

H3K9me2 (1:200) Abcam ab1220, Kimura 6D11 - MABI0307

H3K9me3 (1:200) Kimura 2F3 - MABI0308

H3K27me3 (1:200) Active Motif 61017

H4-pan acetyl (1:500) Active Motif 39925

Pan-histone (1:500) Chemicon/Millipore MAB052

Histone H3 (1:500) Abcam ab1791

FLAG M2 (1:100) Sigma F1804

GFP (1:500) Millipore Sigma MAB3580

GFP (1:500) ThermoFisher Scientific A11122

MET-2 (1:500) Raised against the first 17 amino acids of MET-2 and affinity purified

ARLE-14 (1:500)

Proximity Ligation Assay. Sigma Duolink In Situ Kit (DUO92101) was used for this assay. Embryos were fixed as for regular antibody staining. After overnight staining with primary antibodies at 15oC, the sample was stained at 37oC for 1 hour with secondary antibodies that have oligonucleotide probes attached. Connector oligos were hybridized to the probes and served as templates for circularization by enzymatic ligation when in close proximity. The ligation reaction was incubated at 37oC for 30 minutes. The circularized DNA strands were used for rolling circle amplification (RCA) and the RCA product was detected by hybridizing fluorescently labeled oligos. The RCA reaction was incubated at 37oC for 100 minutes. After each step, slides were washed with TBS + 0.2%

TritonX for 5 minutes. Slides were mounted in DAPI.

Image analysis

Quantitation of histone modifications. Stacks of optical sections were collected with a

ZEISS LSM700 or LSM880 Confocal Microscope and analyzed using Volocity Software.

Nuclei were identified in 3-D using the DAPI channel, and sum signal intensity of histone

72 modification was calculated for each nucleus in interphase. Mitotic nuclei were excluded from the analysis manually based on DAPI morphology. Mean cytoplasmic background was measured at a random point for each embryo, and mean background was subtracted from the nuclear signal by multiplying with nuclear volume. For each nucleus, signal intensity of histone modification was normalized to signal intensity of histones.

Normalized values were averaged for nuclei at given stages and plotted. (Fig. 1D, 2D,

2E, 2J, 3B, 4E, S1D, S3C, S3D).

TEM. Adult hermaphrodites were rapidly chilled in liquid nitrogen while exposed simultaneously to very high pressure (2100 bar). This combination preserves the morphology of organelles and finer structures. Frozen samples underwent freeze substitution to deposit osmium tetroxide for contrast and fixation. Worms were embedded and sectioned along the ovary to view multiple embryos in a row. A detailed protocol is available in (Hall, Hartwieg, and Nguyen 2012).

TEM Image processing. Raw images were processed with Photoshop as 8-bit gray scale images. Images from different preparations were standardized for accurate comparison.

The cytoplasm was used to adjust signal intensity range for each image, but image contrast was not altered. Adjusted images were then saved and quantified with Image J by Line scan analysis.

TEM Line scan analysis: Random lines were drawn across the center of different nuclei, and the intensity measured. Standard deviation was calculated for each individual line.

Standard deviation of 30 lines were averaged for each strain and listed on the plot. A randomly selected line profile for a nucleus is shown as an example. The standard deviation describes the morphology of the nucleus, i.e. a higher standard deviation stems

73 from a more punctate staining pattern that alternates between high and low values (ie electron dense heterochromatin and electron lucent nucleoplasm).

MET-2/ARLE-14 Line scan analysis: Lines that go through the center of the nucleus were drawn across the cell, and the intensity measured. Each line had 100 bins. The intensity in each bin was averaged for 30 lines, and the average line plotted. Error bars denote the standard error of the mean at each bin (Fig. 2I, 4G, S4A, S4F, S4L).

Definition and Quantitation of hubs. Nuclear hubs were defined by intensity thresholding in ImageJ. The threshold was selected manually in wild-type nuclei at the

51-100 cell stage and the same threshold was applied to all the images in a given dataset.

The intensity measurements for each defined hub were averaged to yield the intensity of

“hubs” at given embryonic stages. “Non-hub” was defined as nuclear areas that were below the intensity threshold. The mean intensity in non-hub areas was measured for each nucleus and averaged across 30 nuclei. Non-interphase nuclei were discarded manually. For intensity measurements in the cytosol, at least 4 random areas in the cytosol was chosen for every cell that was in interphase. The measured intensities were averaged.

Half-dose LIN-65 experiments: lin-65 (n3441) moms were crossed with JAC500 his-

72::mCherry males. Progeny of lin-65+/- heterozygotes marked by mCherry were analyzed. On the same slide, SM2333 containing a single copy of zen-4::gfp was used as a wild-type staining control. Mean H3K9me2 intensity in each nucleus was quantified using Volocity and the average H3K9me2 intensity per nucleus plotted, normalized as described in the staining section.

Biochemistry

Harvesting embryos. Mix-staged embryos were collected from adult worms by bleaching

Embryos were, were shaken at 200rpm at 20oC in CSM medium (100mM NaCl, 5.6mM

K2HPO4, 4.4mM KH2PO4, 10μg/ml cholesterol, 10mM potassium citrate, 2mM CaCl2,

2mM MgSO4, 1X trace metals.) without food (Stiernagle 2006). Once the embryos hatched and became L1s, concentrated NA22 bacteria were added to the culture.

Synchronized embryos were harvested by bleaching after 62-66 hours when most worms carried 1-8 embryos, frozen in liquid nitrogen and stored at -80oC.

Immunoprecipitation. Frozen embryo pellets were resuspended in Lysis Buffer (50mM

HEPES pH 7.4, 1mM EGTA, 1mM MgCl2, 100mM KCl, 10% glycerol, 0.05% NP40) with protease inhibitors (Calcbiochem Cocktail Set I, #539131) and incubated on ice for 10 minutes. Using the QSonica Q800 Sonicator, samples were sonicated at 40% amplitude,

10 seconds on, 50 seconds off, for 3 cycles. After sonication, samples were centrifuged for 10 minutes at 10,000g at 4oC. The supernatant was transferred to a new tube and diluted in Dilution Buffer (10mM Tris-Cl pH 7.5, 150mM NaCl, 0.5mM EDTA). 1.5 mg of total lysate was pre-cleared for 1 hour at 4oC with 25 μl Chromotek magnetic agarose beads prior to IP. The pre-cleared lysate was then used for immunoprecipitating MET-

2::GFP With Chromotek GFP-TRAP magnetic agarose beads or Sigma FLAG M2 antibody coupled to magnetic agarose beads for 5 hours at 4oC. Beads were rinsed with

Dilution Buffer 3 times, washed with Dilution Buffer twice for 5 minutes, and eluted with

50μl 0.2M glycine pH 2.5 for 30 seconds under constant mixing or with 200μg/ml 3xFLAG peptide in TBS for 1 hour at 4oC. 5μl 1M Tris base pH 10.4 was added for neutralization after glycine elution. Samples were boiled in 2X Laemmli Sample Buffer (Biorad #161-

0737) with 50mM DTT and analyzed by Western blotting or silver staining (SilverQuest

Silver Staining Kit, ThermoFisher, LC6070). The FLAG M2 antibody was coupled to

Pierce Protein A/G magnetic beads (#88802) using the Pierce Crosslink Magnetic IP/Co-

IP Kit (#88805).

Mass spectrometry. Reagents and Chemicals: Deionized water (18.2 M, Barnstead,

Dubuque, IA) was used for all preparations. Buffer A consists of 5% acetonitrile 0.1% formic acid, buffer B consists of 80% acetonitrile 0.1% formic acid, and buffer C consists of 500 mM ammonium acetate and 5% acetonitrile.

Sample Preparation: Proteins were precipitated in 23% TCA (Sigma-Aldrich, St. Louis,

MO, Product number T-0699) at 4 °C O/N. After 30 min centrifugation at 18000 x g, protein pellets were washed 2 times with 500 ul ice-cold acetone. Air-dried pellets were dissolved in 8 M urea/ 100 mM Tris pH 8.5. Proteins were reduced with 1 M Tris(2- carboxyethyl)phosphine hydrochloride (Sigma-Aldrich, St. Louis, MO, product C4706) and alkylated with 500 mM 2-Chloroacetamide (Sigma-Aldrich, St. Louis, MO, product

22790-250G-F). Proteins were digested for 18 hr at 37 °C in 2 M urea, 100 mM Tris pH

8.5, 1 mM CaCl2 with 2 ug trypsin (Promega, Madison, WI, product V5111). Digestion was stopped with formic acid, 5% final concentration. Debris was removed by centrifugation, 30 min 18000 x g.

MudPIT Microcolumn: A MudPIT microcolumn (Dirk A. Wolters, Michael P. Washburn, and John R. Yates 2001) was prepared by first creating a Kasil frit at one end of an undeactivated 250 m ID/360 m OD capillary (Agilent Technologies, Inc., Santa Clara,

CA). The Kasil frit was prepared by briefly dipping a 20 - 30 cm capillary in well-mixed

300 L Kasil Kasil 1624 (PQ Corporation, Malvern, PA) and 100 L formamide, curing at

100OC for 4 hrs, and cutting the frit to ~2 mm in length. Strong cation exchange particles

(SCX Partisphere, 5 m dia., 125 Å pores, Phenomenex, Torrance, CA) was packed in- house from particle slurries in methanol 2.5 cm. Additional 2.5 cm reversed phase particles (C18 Aqua, 3 µm dia., 125 Å pores, Phenomenex) were then similarly packed into the capillary using the same method as SCX loading, to create a biphasic column.

An analytical RPLC column was generated by pulling a 100 m ID/360 m OD capillary

(Polymicro Technologies, Inc, Phoenix, AZ) to 5 m ID tip. Reversed phase particles

(Aqua C18, 3 m dia., 125 Å pores, Phenomenex, Torrance, CA) were packed directly into the pulled column at 800 psi until 12 cm long. The MudPIT microcolumn was connected to an analytical column using a zero-dead volume union (Upchurch Scientific

(IDEX Health & Science), P-720-01, Oak Harbor, WA). LC-MS/MS analysis was performed using an Agilent Technologies 1200 HPLC pump and a Thermo Orbitrap Velos using an in-house built electrospray stage. MudPIT experiments were performed with steps of 0% buffer C, 30% buffer C, 50% buffer C, 90/10 % buffer C/B and 100% C, being run for 3 min at the beginning of each gradient of buffer B. Electrospray was performed directly from the analytical column by applying the ESI voltage at a tee (150

m ID, Upchurch Scientific)(Dirk A. Wolters, Michael P. Washburn, and John R. Yates

2001). Electrospray directly from the LC column was done at 2.5 kV with an inlet capillary temperature of 325 OC. Data-dependent acquisition of MS/MS spectra with the Orbitrap

Velos were performed with the following settings: MS/MS on the 10 most intense ions per precursor scan; 1 microscan; reject unassigned charge state and charge state 1; dynamic

77 exclusion repeat count, 1; repeat duration, 30 second; exclusion list size 200; and exclusion duration, 30 second.

Data Analysis: Protein and peptide identification and protein quantitation were done with

Integrated Proteomics Pipeline - IP2 (Integrated Proteomics Applications, Inc., San

Diego, CA. http://www.integratedproteomics.com/). Tandem mass spectra were extracted from raw files using RawConverter (He et al. 2015) with monoisotopic peak option and were searched against Wormbase protein database (WB257) with reversed sequences using ProLuCID (Junmin Peng et al. 2002; “ProLuCID: An Improved

SEQUEST-like Algorithm with Enhanced Sensitivity and Specificity” 2015). The search space included all half and fully-tryptic peptide candidates. Carbamidomethylation

(+57.02146) of cysteine was considered as a static modification. Peptide candidates were filtered using DTASelect with the parameters -p 2 -y 1 --trypstat --pfp 0.01 --extra --pI -

DM 10 --DB --dm -in -m 1 -t 2 --brief --quiet (Tabb, McDonald, and Yates 2002). Mass spectrometry identified a total of 602 proteins in the MET-2::GFP sample. 500 proteins were eliminated due to their presence in controls, and 43 proteins were eliminated because they had low sequence coverage (<6%). Remaining 59 proteins were further analyzed based on suggested function and localization from literature. Promising candidates were chosen for a loss-of-function screen.

RNA expression analysis. Embryos frozen in liquid nitrogen were partially thawed. An equal volume of glass beads (Sigma G8772) were added and samples were vortexed for a minute. For Trizol-chloroform extraction, 1ml Trizol (Thermo Fischer Scientific

#15596026) was added to the sample, vortexed for 30 seconds and incubated at room temperature for 5 minutes. 300 μl chloroform (Calbiochem, Omnipur #3155) was added,

78 shaken by hand and incubated at room temperature for 3 minutes. The sample was centrifuged at 14000rpm at 4oC for 15 minutes. Upper aqueous phase was transferred to a fresh tube and RNA was precipitated using 500μl isopropanol and 1 μl Glycoblue

(Thermo Fisher Scientific AM9516). The pellet was air-dried and resuspended in nuclease-free water at room temperature for 10 minutes. Samples were treated with

DNAse at 37oC for 30 minutes using the Turbo DNA-free kit (Thermo Fisher Scientific

AM1907), and 500 ng of RNA was used for reverse transcription with a random primer mix (Protoscript First Strand cDNA Synthesis Kit, NEB E6300). The synthesis reaction was diluted in water to yield a total volume of 50μl, and 3μl of the cDNA was analyzed by qPCR (KAPA SYBR FAST qPCR Master Mix, KK4601).

ChIP. H3K9me2 ChIP was done as described previously (Hsu et al. 2015) with the following changes: Embryos frozen in liquid nitrogen were thawed on ice and fixed in

1.5% Formaldehyde (Electron Microscopy Sciences, #15686) for 15 minutes at room temperature. Using the QSonica Q800 Sonicator, samples were sonicated at 30% amplitude, 30 seconds ON, 30 seconds OFF for a total of 15 minutes at 4oC, yielding 100-

300 base pair fragments. 6μl of Kimura 6D11 antibody was coupled to 25μl beads (Pierce

Protein A/G magnetic beads, #88802) for 8 hours at 4oC prior to ChIP. 40 μg of chromatin was used per ChIP reaction and chromatin was pre-cleared for 2 hours at 4°C using uncoupled magnetic beads (Pierce #88802). To elute the bound immunocomplexes, 150

μL of elution buffer (50mM NaHCO3, 140mM NaCl, 1% SDS) was added to each tube and heated at 65°C for 15 minutes. For MET-2::GFP ChIP, embryos were fixed with

1,5mM EGS (Pierce #21565) for 10 minutes and with 1% Formaldehyde for another 10 minutes. 6μl of GFP antibody (Abcam5665) was coupled to 25μl beads for 8 hours at 4oC

79 prior to ChIP. 2μl of H3K4me3 (Abcam8580) antibody was coupled to 25μl beads for 8 hours at 4oC prior to ChIP.

Library preparation: The ChIP-Seq libraries were generated by using Apollo 324 System and PrepX ILM DNA Library Kit from IntergenX. After adaptor ligation, the input and ChIP

DNA were enriched by PCR amplification using NEBNext High-Fidelity 2X PCR master

Mix with Q5 polymerase and PrepX PCR primer with the following PCR conditions: 30 seconds at 98°C, [10 seconds at 98°C, 30 seconds at 60°C, 30 seconds at 72°C] for 8 cycles for Input libraries and 11 cycles for ChIP libraries, following 5 minutes at 72°C

(~15 uL adaptor ligated DNA, 25 uL NEBNext High-Fidelity 2X PCR master Mix, 2uL

Universal PCR primer, brought to to 50ul with Nuclease-Free water). The enriched DNA was then purified using 50 uL (1:1 ratio of DNA volume and beads) of PCR Clean DX

Beads (Aline) and size selected by Pippin Prep, 180-600bp. 1 uL of each library was applied to measure the concentration using a Qubit dsDNA assay kit (Invitrogen). 1 ng of

DNA from each library was checked by a TapeStation (Agilent Technologies). Input and

ChIP libraries were pooled such that they each had the same amount of molecules and expected for obtaining the similar number of reads. The Illumina sequencing was performed with 75 nt paired-end reads.

Sequencing Analysis: DNA fragments were sequenced on an Illumina HiSeq machine, yielding 64-95 million 75bp paired-end reads per sample. Reads were aligned to the C. elegans reference genome (ce10) with Bowtie2 (Langmead and Salzberg 2012), version

2.3.1, and default parameters except for ‘--very-sensitive’ and ‘-X 2000’. PCR duplicates were removed from the alignment files after identifying properly paired fragments that shared both leftmost and rightmost genomic coordinates. MACS2 (Zhang et al. 2008),

80 version 2.1.1.20160309, was used to call peaks in the ChIP samples, using input DNA as the control and analyzing only properly paired fragments (-f BAMPE). Peaks were annotated using ChIPseeker (Yu, Wang, and He 2015), version 1.12.0. Log-likelihood ratio tracks were calculated using the MACS2 module ‘bdgcmp’, and correlations were calculated using the BigWig tools ‘bedGraphToBigWig’ and ‘wigCorrelate’ (Kent et al.

2010).

Data access. The data from this study will be submitted to NIH SRA and GEO database with accession numbers (pending).

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CHAPTER 3

ONSET OF HETEROCHROMATIN FORMATION IS DICTATED

BY TIME AFTER FERTILIZATION

3.1 Abstract

Changes in chromatin organization accompany embryonic development, but it is poorly understood how the timing of these changes are set. During the first hour of development,

C. elegans embryos exhibit a surge in di-methylated histone H3 Lysine 9 (H3K9me2), a repressive histone modification that initiates the onset of heterochromatin. Here we test whether classical embryonic time-keeping mechanisms dictate the timing of H3K9me2.

Surprisingly, timing of H3K9me2 establishment does not rely on mechanisms that count cells or nuclei, does not require Polymerase II transcription and is not dependent on RNA interference. Instead, the onset of H3K9me2 tracks with time after fertilization. The cumulative length of interphase during early embryogenesis is important for the nuclear accumulation and activity of the H3K9me2 methyltransferase MET-2 and its binding partners. These results reveal the key determinants that set the timing of H3K9me2 establishment during embryogenesis.

3.2 Introduction

Embryogenesis is a multi-faceted process, where cellular differentiation and molecular changes are coordinated in time and space to produce coherent development. While a lot is known about the molecular components involved, the temporal regulation of these events is not well understood. A molecular hallmark of differentiation is the segregation of the genome into euchromatin and heterochromatin domains to regulate gene expression (Fadloun, Eid, and Torres-Padilla 2013; Politz, Scalzo, and Groudine 2013).

Heterochromatin is characterized by a high frequency of repetitive sequences and condensed chromatin that is rich in silencing marks such as H3K9 methylation and its ligand HP1 (Lachner et al. 2001; Rice et al. 2003; Rübe et al. 2011; Yuan and O’Farrell

2016).

C. elegans embryos form heterochromatin during the first hour of embryogenesis, which can be observed with three assays: First, analyzing the morphology of exogenous, artificial chromosomes has revealed that chromatin compacts as embryos develop

(Yuzyuk et al. 2009). Second, ultrastructural analysis of the nucleus by electron microsopy has shown emergence of electron-dense heterochromatin domains during gastrulation (Chapter 2). Third, H3K9 di-methylation, a repressive histone modification, is low in early embryos and rises during gastrulation (Chapter 2). The H3K9me1/me2 methyltransferase met-2 is required for all three processes (Chapter 2, (Fakhouri et al.

2010; Towbin et al. 2012)), revealing that H3K9 di-methylation is critical for heterochromatin formation during embryogenesis.

RNAi pathways are critical to target heterochromatin machinery in worms and other organisms such as S. pombe (Moazed 2009). In particular, the piwi pathway and the nuclear RNAi (nrde) pathway both target repeat sequences that make up the bulk of heterochromatin and are marked by H3K9me2 (McMurchy et al. 2017; Ashe et al. 2012;

Weick and Miska 2014). While it is known that the RNAi machinery targets H3K9 methylation, it is not clear whether these components dictate the timing of H3K9me2 in embryos.

There are several models for embryonic time-keeping which are not mutually exclusive and could be applied to H3K9 methylation. The first model postulates that cell counting mechanisms such as the amount of DNA in the embryo could be a timer

(Newport and Kirschner 1982b; Dekens et al. 2003). The second model focuses on the changes in the nuclear-to-cytoplasmic ratio as embryonic cells divide, which could be critical for diluting or concentrating a maternal repressor/activator (Newport and Kirschner

1982a; Pritchard and Schubiger 1996). The third model predicts that a signaling cascade is initiated at fertilization and acts as an absolute clock by regulating cell cycle complexes

(Kimelman, Kirschner, and Scherson 1987; Howe and Newport 1996; Ferree, Deneke, and Di Talia 2016; Yuan et al. 2016; Treen et al. 2018). A fourth possibility is that initiation of zygotic transcription is involved in driving chromatin re-organization (Tadros and

Lipshitz 2009). It remains to be tested whether any of these models dictate the timing of

H3K9me2 during embryogenesis.

Here we systematically test the molecular determinants for timing the establishment of H3K9me2 and rule out Polymerase II transcription, RNAi machinery or cell counting mechanisms as potential timers. Instead, timing of H3K9me2 is dictated by

91 an absolute clock. Slowing down early embryonic cell divisions leads to precocious nuclear accumulation and activity of MET-2 and its binding partners, which is critical for

H3K9 methylation.

3.3 Results

H3K9 de-methylation is not a timer for establishing H3K9me2 domains at gastrulation.

We began our analysis by testing different models to understand what dictates the timing of H3K9 methylation during embryogenesis. We focused on the earliest stages of embryogenesis that are critical for setting up heterochromatin domains (Chapter 2, Figure

3.1A). Many changes occur as early embryonic cells divide, and the initiation of gastrulation around the 28-cell stage is a developmental milestone (Figure 3.1A). In pre- gastrula embryos, MET-2 is mostly cytosolic and gradually translocates into the nucleus to methylate histone H3 at gastrulation (Figure 3.2A-B). We previously demonstrated that the regulation of MET-2 by two binding partners LIN-65 and ARLE-14 is crucial for initiating H3K9 methylation (Chapter 2). Here, we explore additional time-keeping mechanisms in the embryo that could act on MET-2, LIN-65 and ARLE-14 or regulate

H3K9 methylation.

Initially, we wondered if an H3K9me2 demethylase might act in early embryos to remove H3K9me2. To test this idea, we examined a mutant for the H3K9me2 demethylase jmjd-1.2 (Kleine-Kohlbrecher et al. 2010). In jmjd-1.2(tm3713) mutants that don’t produce JMJD-1.2 (Myers et al. 2018), H3K9me2 was still extremely low in early embryos and was established normally at gastrulation (Figure 3.1C-E), suggesting that

92 histone de-methylation is not part the H3K9me2 timer, and focusing our attention on MET-

2 and its partners.

Figure 3.1. H3K9me1/me2 methyltransferase MET-2 is regulated during embryogenesis to form heterochromatin. A. Diagram showing a timeline for C. elegans embryogenesis and a summary of changes that accompany differentiation. With each cell division and passing time, the total number

Figure 3.1 continued of cells, amount of DNA per embryo and the nuclear-to-cytoplasmic ratio per cell increases. The transition from light to dark green represents initiation of gastrulation around the 28-cell stage. Zygotic transcription is initiated at the 4-cell stage, but a big wave coincides with gastrulation (Schauer and Wood 1990; Storfer-Glazer and Wood 1994; Levin et al. 2012). All of these changes are candidates for dictating the timing of H3K9me2. B. MET-2::GFP localization and H3K9me2 levels during embryogenesis. MET-2::GFP (green), H3K9me2 (red), DAPI (blue), Scale bar 2 μm. Note that MET-2::GFP accumulates in the nucleus as H3K9me2 domains are formed. C. H3K9me2 levels in wild-type vs. H3K9me2 demethylase jmjd-1.2 mutants at pre- gastrula (4-cell and 8-cell stage) and gastrula (21-50 cell) stages. D-E. Single nuclei in interphase showing H3K9me2 levels in wild-type vs. jmjd-1.2 mutants. Quantitation of H3K9me2 levels per nucleus. Early jmjd-1.2 embryos do not have increased H3K9me2.

Zygotic transcription is not rate-limiting for H3K9 di-methylation.

The onset of gastrulation and surge in H3K9me2 is accompanied by a big wave in zygotic transcription (Edgar, Wolf, and Wood 1994; Storfer-Glazer and Wood 1994; Baugh 2003;

Yuzyuk et al. 2009; Levin et al. 2012; Hsu et al. 2015), raising the possibility that it’s the rate-limiting step for the deposition of H3K9me2. We hypothesized that met-2 or its positive regulators could become activated in the embryo as zygotic transcription begins

(Figure 3.2A, Model 1). Alternatively, transcription elongation could be rate-limiting for recruiting chromatin modifiers like MET-2 to specific loci through interactions with the

RNAi machinery ((Guang et al. 2010), Figure 3.2A, Model 2). To test this idea, we blocked zygotic transcription and examined whether the onset of H3K9me2 was disturbed. Our experimental design enabled us to compare H3K9me2 levels across different genotypes accurately: In each experiment, we included wild-type embryos marked with a single copy of ZEN-4::GFP or HIS-72::mCHERRY as an on-slide control and processed wild-type and mutant embryos together on the same slide (Figure 3.2B).

To block transcription, we inactivated the canonical TFIID subunit, taf-6.2, by a temperature sensitive mutation (ax514) (Bowman, Seydoux, and Kelly 2011), and a core subunit of RNA Polymerase II, ama-1, by RNAi. To assess the strength of the block on transcription, we stained embryos with the H5 Polymerase II antibody against the phosphorylated Ser2 on the CTD domain, which is a hallmark of transcriptional elongation. Under our conditions, H5 Polymerase II signal was undetectable after blocking zygotic transcription, whereas non-specific staining of P-granules by the H5 antibody served as a positive control. (Figure 3.2C, D). Despite the lack of detectable H5,

H3K9me2 levels were identical to wild-type embryos (Figure 3.2C, E), suggesting that zygotic genes or elongating Polymerase II are not rate-limiting for H3K9me2 onset.

The transcriptional block suggested that MET-2 and its regulators must be contributed by the mother and be independent of zygotic transcription. To test this idea, we looked at the expression of met-2 and its binding partners lin-65 and are-14, which are required for H3K9me2. met-2, lin-65 and arle-14 transcripts were abundant in early embryos before the initiation of zygotic transcription (Levin et al. 2012; Tintori et al. 2016) revealing that these RNAs were deposited by the mother. To test if zygotic MET-2 was required for embryonic H3K9me2, we mated met-2 mutant mothers with wild-type males and analyzed the progeny, which lacked maternal MET-2 but contained a zygotic copy of the met-2 locus. We found that zygotic met-2 could not rescue H3K9me2 in embryos, and

H3K9me2 was undetectable in the absence of maternal MET-2 (Figure 3.2F). Similarly, lin-65 and arle-14 mutants exhibit significantly reduced H3K9me2 levels, and neither zygotic lin-65 nor arle-14 could rescue H3K9me2 (Figure 3.2G). Moreover, when we mated wild-type mothers with met-2::gfp males, the resulting met-2::gfp progeny never

95 expressed GFP (Figure 3.2H,I). This result demonstrates that maternally deposited factors are responsible for establishing H3K9me2 in embryos, and that the onset of transcription cannot account for the timing of H3K9me2 deposition.

Figure 3.2. Zygotic transcription is not rate-limiting for H3K9 di-methylation. A. Two models describe how initiation of zygotic transcription might be important for dictating the timing of H3K9me2. Model 1: Transcription could be important for the Figure

Figure 3.2 continued production of MET-2 protein or an activator of MET-2. Model 2: Transcription itself might be rate-limiting to recruit MET-2 to chromatin (Bühler and Moazed 2007; Guang et al. 2010) B. Experimental design. Wild-type embryos (green) containing a single copy ZEN-4::GFP or HIS-72::mCHERRY marker were analyzed on the same slide as unlabeled mutant embryos (yellow). This approach controlled for staining variability, thereby enabling an accurate comparison between different genotypes. C. Gastrula embryos stained with an antibody that detects the elongating form of RNA Polymerase II (H5; green) or H3K9me2 (red). Scale bar 2 μm. Transcription was blocked by combining ama-1 RNAi with the ax514 mutation in the initiation factor taf-6.2. White arrow points to non-specific P-granule (Wang and Seydoux 2014) staining by the H5 antibody and serves as a staining control. D-E. Quantitation of mean signal intensity per nucleus for Pol II and H3K9me2 in wild- type vs. transcriptionally impaired embryos on the same slide. Note that transcription elongation is undetectable but H3K9me2 levels are unaffected. F-G. H3K9me2 levels in gastrula embryos that lack maternal met-2, lin-65 or arle-14 but contain a paternal copy. The genotype of the mother is highlighted in red, the father’s in blue. Zygotic copies of these genes cannot rescue H3K9me2 in the embryo. H-I. MET-2::GFP levels in gastrula embryos that lack a maternal copy of the construct, but inherit a paternal copy. Note that the paternal copy of MET-2 is not expressed in the embryo and maternal MET-2 sufficient for all H3K9me2 during embryogenesis.

RNAi machinery components do not dictate the timing of H3K9me2.

The small RNA pathway engages with the nrde pathway in the nucleus to recruit histone methyltransferases during transcriptional elongation (Guang et al. 2010; Weick and Miska

2014). MET-2 synergizes with the small RNA pathways to silence repetitive sequences, which are enriched for H3K9me2 (McMurchy et al. 2017). On the other hand, the independence of H3K9me2 from Pol II transcription suggested that the RNAi machinery might not be rate-limiting during embryogenesis to target MET-2 to transcribing genes.

Consistent with this idea, we observed that a null allele of the Piwi Argonaute prg-1 (Wang and Reinke 2008) had no effect on H3K9me2 accumulation (p=0.44), nor did mutations in the nuclear argonaute nrde-3 (p=0.21) (Figure 3.3A, B) (Guang et al. 2008). nrde-2 mutants (Guang et al. 2010) exhibited a subtle reduction in H3K9me2, but the effect was

97 not significant (p=0.07). (Figure 3.3B). We conclude that these factors may help target

H3K9me2 to certain loci, but they are not essential for the timely acquisition of H3K9me2 in embryos.

We also tested the RNAi resistant mago6 strain which carries six inactivated argonaute genes: sago-1, sago-2, ppw-1, F58G1.1 and predicted pseudogenes C06A1.4 and M03D4.6 (Yigit et al. 2006). In the mago6 strain, H3K9me2 was established at gastrulation, like the wild type (Figure 3.3C-D), revealing that RNAi components do not dictate the timing of H3K9me2.

Figure 3.3. RNAi machinery is not the timer for H3K9me2. A. H3K9me2 staining in wild-type embryos containing a single copy ZEN-4::GFP vs. prg- 1 (piwi) and nrde-2 or nrde-3 (nuclear RNAi) mutants. Note that the wild-type embryos were located on the same slide as the mutants, as a staining control. Note that H3K9me2 is still established at gastrulation in RNAi mutants. B. H3K9me2 signal intensity normalized to histone H3 for embryonic nuclei at the gastrula stage in wild-type vs. mutant embryos. C. H3K9me2 staining in wild-type embryos containing a single copy HIS-72::mCHERRY and mago6 mutant embryos containing mutations in 6 argonautes: sago-2, ppw-1, C06A1.4, F58G1.1,M03D4.6, sago-1 on the same slide. D. Quantitation of mean signal intensity per nucleus for H3K9me2 in wild-type vs. mago6 mutant embryos. H3K9me2 levels are not altered in mago6 mutants.

Length of interphase, but not cell counting mechanisms, dictates timing of

H3K9me2.

The transcription block and RNAi survey ruled out some expected mechanisms for

H3K9me2 onset. To gain a broader perspective on regulation of heterochromatin onset, we considered whether early embryonic cell divisions contribute to timing. Classical models in developmental biology (Tadros and Lipshitz 2009) postulate that DNA synthesis and mitosis enable cells to keep time by titrating a DNA-associated repressor, or reaching a critical nuclear-to-cytoplasmic ratio. We wondered whether a similar process might regulate the timing of H3K9me2 deposition in C. elegans.

To determine if cell counting is important, we uncoupled the number of cells produced over a given unit of time. We inactivated div-1, a subunit of DNA polymerase- alpha primase complex, by using a temperature-sensitive mutation (or148) to extend the duration of S-phase and slow down cell divisions (Encalada et al. 2000). One hour after the 2-cell stage, wild-type embryos had 25-30 cells, whereas div-1 mutant embryos had

5-15 cells (Figure 3.4A). div-1 embryos exhibited precocious H3K9me2 based on cell number, but wild-type levels based on time post-fertilization (Figure 3.4B, C). The amount of H3K9me2 per nucleus was similar in 25-30 cell wild-type embryos and 5-15 cell div-1 embryos. These delayed div-1 cells had a similar volume to wild-type embryos with the equivalent number of cells (Figure 3.4D) and comparable amounts of DNA as wild-type embryos that contain the same number of cells (Figure 3.4E). This result rules out some counting models, specifically the nuclear-to-cytoplasmic ratio, the number of cells or the amount of DNA. In short, it is possible to acquire high levels of H3K9me2 without reaching a certain cell number, nuclear-to-cytoplasmic ratio or undergoing a certain number of cell

Figure 3.4. Slowing down S-phase leads to precocious accumulation of H3K9me2 and its regulators. A. Time-line of embryogenesis with respect to absolute amount of time and number of cells per embryo in wild-type (green) or div-1(or148) mutant (gray) embryos.

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Figure 3.4 continued B-C. H3K9me2 staining in embryos that contain the same number of cells (8-cell, left panels) or that have developed for the same amount of time after the 2-cell stage (1 hour, right panels). Scale bar 2 μm. Amount of H3K9me2 tracks with absolue time, not number of cells. D. Nuclear volume in wild-type vs. div-1(or148) embryos at the 8-cell stage. E. Mean DAPI intensity in wild-type vs. div-1(or148) embryos at the 8-cell stage. F,I. Wild-type vs div-1 (or148) embryos at the pre-gastrula stage stained with an antibody against endogenous MET-2 on the same slide. Quantitation of protein levels per nucleus. G,J. Progeny of lin-65::3xflag worms fed with empty vector vs div-1 RNAi. Embryos at the pre-gastrula stage were stained with a FLAG antibody. Quantitation of protein levels per nucleus. H,K. Wild-type vs div-1 (or148) embryos at the pre-gastrula stage stained with an antibody against endogenous ARLE-14 on the same slide. Quantitation of protein levels per nucleus. divisions. Instead, this result suggests that the timing of H3K9me2 is dictated by an absolute clock (~1 hour after the 2-cell stage). div-1 mutants specifically extend S-phase compared to wild-type embryos (Encalada et al. 2000), revealing that the cumulative time early embryonic cells spend in interphase is critical for measuring time.

Nuclear MET-2 is rate-limiting for H3K9me2 levels and accumulates in the nucleus together with binding partners LIN-65 and ARLE-14 over time (Chapter 2). We hypothesized that slowing cell divisions in early div-1 mutant embryos might allow these factors to accumulate in the nucleus more readily. A prediction of this hypothesis would be more nuclear MET-2 in div-1 embryos with precocious H3K9me2 compared to wild- type. Indeed, MET-2 and its binding partners each accumulated in the nucleus earlier compared to wild-type embryos (Figure 3.4F-K).

One concern was that the replicative stress or DNA damage in early div-1 embryos caused higher levels of H3K9 methylation to engage DNA repair pathways (Ayrapetov et al. 2014). We hypothesized that if DNA damage in div-1 mutants caused the increase in

H3K9me2, one might expect restoring the faster cell cycle in div-1 mutants to lead to even

101 more DNA damage and H3K9me2 (Figure 3.5A). Alternatively, if the amount of time in interphase was the critical parameter, then restoring the faster cell cycle to div-1 mutants would restore/rescue normal timing of H3K9me2 accumulation. To restore faster cell cycle progression to div-1 mutant embryos, we inactivated the ATR related gene atl-1 which leads to faster cell cycles but potentially more DNA damage (Brauchle, Baumer, and Gönczy 2003). atl-1(RNAi); div-1 double mutants partially suppressed the increase in H3K9me2 (Figure 3.5B-D), suggesting precocious H3K9me2 in div-1 mutants was not due to DNA damage. Instead, timing of H3K9me2 depends on the amount of time from fertilization. We hypothesize that the cumulative time spent in S-phase/interphase after fertilization permits re-localization and activation of MET-2, and thereby dictates the onset of H3K9me2.

Figure 3.5

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Figure 3.5 continued. Precocious H3K9me2 in div-1 mutants is not due to DNA damage, but depends on amount of time spent after fertilization. A. Rationale. Increased H3K9me2 in div-1 mutants could be due to I) the extension of S- phase (light orange) or II) DNA damage (dark orange). Accordingly, restoring the cell cycle by inactivating atl-1 in div-1 mutants is predicted to have two alternative outcomes: I) rescue of cell cycle timing and reduction in H3K9me2 (light orange) or II) additional DNA damage and increased H3K9me2 (dark orange). B. H3K9me2 staining in wild-type vs. atl-1 RNAi, div-1(or148) and atl-1 RNAi; div-1(or148) double inactivation embryos at the pre-gastrula stage. Color code: wild-type – green, atl- 1 – purple, div-1 – gray, atl-1;div-1 – blue. In each experiment, wild-type embryos containing zen-4::gfp were included as on-slide controls (not shown). C-D. Single nuclei in interphase showing H3K9me2 levels and quantitation. Note that atl- 1 RNAi partially rescues H3K9me2 levels in div-1 mutants in support of Scenario I (light orange from A).

MET-2 promotes loss of plasticity.

The dynamics of heterochromatin are important for proper development of other organisms (Fadloun, Eid, and Torres-Padilla 2013) but have not been examined in C. elegans. In worms, H3K9me2 appears when gastrulating embryos lose developmental potential (Chapter 2), raising the possibility that MET-2 is required to restrict cell fate.

Other chromatin modulators such as mes-2/Enhancer of zeste and glp-1/Notch are known to contribute to loss of embryonic plasticity (Djabrayan et al. 2012; Yuzyuk et al. 2009), and we took advantage of the Cell Fate Challenge Assay (Horner et al. 1998; Mango

2009) to test the importance of met-2 in loss of plasticity (Figure 3.6A). Briefly, embryos were challenged to alter their development and acquire muscle fate by ectopic expression of hlh-1/MyoD under control of the heat-shock promoter (HS::hlh-1; (Fukushige and

Krause 2005)). We chose the 80-100-cell stage to measure plasticity because H3K9me2 was very different in wild-type vs. mutant embryos at that stage (Chapter 2). In addition, the majority of wild-type embryos lose plasticity by the 80-100 cell stage (Mango 2009).

The foregut marker PHA-4 was used to identify cells that retained their endogenous

103 identity and resisted exogenous HLH-1 (Horner et al. 1998; Yuzyuk et al. 2009). These markers for differentiation reflected endogenous genes to ensure that we tracked native expression. Exogenous reporters can sometimes lead to inaccurate expression (Mango

2007). As a control, we examined hlh-1 mRNA expression before and after heat shock and observed no difference in induction between wild-type and mutant embryos (Figure

3.6D).

When challenged with HS::hlh-1, 35% of 100-cell-stage wild-type embryos completely converted to muscle fate. In met-2 mutants, 54% of embryos responded to

HS::hlh-1 with a complete cell-fate transformation (Figure 3.6C, 100-cell stage, n=4, >100 embryos each, p=0.008). This result suggests that, normally, met-2 promotes loss of plasticity during early gastrulation. By the ~200 cell stage, none of the wild-type or met-2 mutant embryos remained fully plastic (Figure 3.6C). Moreover, both wild-type and met-

2 mutant embryos had a similar number of cells that resisted changing fate, suggesting met-2 mutants eventually terminate plasticity (data not shown). The data indicate that met-2 restricts developmental plasticity during gastrulation but is not absolutely required.

H3K9me3 is deposited by SET-25, and partially depends on H3K9me2 by MET-2

(Towbin et al. 2012). We wondered if SET-25 was also important for terminating plasticity.

Surprisingly, inactivation of set-25 lead to the opposite result: only 20% of set-25 embryos were developmentally plastic (n=3 experiments, >60 embryos each, p=0.011). This result demonstrated that H3K9me3 was not required to terminate plasticity during gastrulation.

Instead, developmental plasticity correlated with level of H3K9me2, which is low in met-2 and high in set-25 mutants relative to wild-type (Chen et al., in review). To test the importance of H3K9me2, we examined met-2; set-25 double mutants, which lack

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Figure 3.6. MET-2 promotes loss of developmental plasticity at mid-gastrulation. A. Cell Fate Challenge Assay (CFCA). Muscle fate regulator hlh-1/MyoD is induced in somatic cells. Terminally differentiated embryos stained for induced or endogenous fate markers (muscle paramyosin, red, or foregut PHA-4, green, respectively (Fukushige and Krause 2005; Horner et al. 1998)). B. Terminally differentiated embryos. Arrowheads depict cells lacking paramyosin, some of which express PHA-4. Scale bar, 5µm. C. Percentage of wild-type (n=109) vs. met-2 (n=136) or wild-type (n=64) vs. set-25 (84) or wild-type (n=31) vs. met-2; set-25 (n=38) embryos scored as developmentally plastic based on absent PHA-4 and ubiquitous paramyosin. Note that MET-2 promotes loss of plasticity, while SET-25 prevents it.

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Figure 3.6 continued D. hlh-1/MyoD mRNA levels normalized to eft-3 in wild-type or mutant embryos, with (+HS) or without (-HS) heat-shock induction. CFCA results are not due to differential induction of hlh-1 in wild-type vs. mutants. E. GO Term Analysis for genes enriched for H3K9 methylation. Genes associated with H3K9me2 are involved in regulating plasticity, whereas H3K9me1 and H3K9me3 are not.

H3K9me2 (Towbin et al. 2012; Garrigues et al. 2014). Double mutants had prolonged plasticity (Figure 3.6C, n=2, 30 embryos each, p=0.045), revealing that the reduced plasticity of set-25 mutants required met-2 activity. A simple hypothesis is that H3K9me2 modulates developmental plasticity. Among genes with peaks of H3K9me2, there was enrichment for GO terms “cell fate determination” and “embryonic pattern specification,” which included genes expressed in the pre-gastrula embryo (e.g. par-1, par-4, mbk-2 and others). These GO terms were not enriched for H3K9me1 or H3K9me3, likely explaining their distinct roles in the embryo (Figure 3.6E).

3.4 Discussion

This study has made three contributions to understand the timing of H3K9me2 during embryogenesis. First, it has ruled out several models including cell counting, nuclear-to- cytoplasmic ratio, zygotic transcription and RNAi machinery as timers. Second, it has revealed that the absolute amount of time early embryonic cells spend in S-phase determines the onset of H3K9 di-methylation. Third, we have shown that the H3K9me2 methyltransferase MET-2 promotes the timely loss of developmental plasticity at gastrulation.

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Zygotic Transcription and RNAi machinery are not timers.

An intriguing model is that the absence of H3K9 methylation in early embryos provides a window of opportunity where repetitive sequences can be transcribed and be targeted by the RNAi machinery to acquire H3K9me (Yu et al. 2013; Penke et al. 2016; Zeller et al.

2016; McMurchy et al. 2017).

The Argonaute (Ago) family of proteins and the small RNAs that program them are central players in RNA silencing and heterochromatin assembly (Moazed 2009). In worms, both exogenous small RNAs and endogenous piRNAs converge on the WAGO secondary siRNA pathway, which serves as a critical amplification step before silencing

(Pak and Fire 2007; Gu et al. 2012). In the cytosol, the exo-RNAi argonaute RDE-1 or the piwi argonaute PRG-1 triggers production of secondary siRNAs (Yigit et al. 2006; Ashe et al. 2012; Weick and Miska 2014). Upon binding to the secondary siRNAs, nuclear RNAi

(nrde) argonaute NRDE-3 translocates from the cytosol to the nucleus to promote H3K9 methylation (Guang et al. 2008, 2010; Burkhart et al. 2011; Burton, Burkhart, and

Kennedy 2011).

In our studies, we focused on H3K9me2, as it is thought to precede H3K9me3 and increases more dramatically compared to H3K9me3 during gastrulation (Towbin et al.

2012). Under our conditions, piwi Argonaute PRG-1, nrde Argonautes NRDE-2 and

NRDE-3, or six WAGO argonautes were not rate-limiting for H3K9me2 in the embryo

(Figure 3.3).

The nrde pathway acts co-transcriptionally and inhibits Polymerase II during the elongation phase of transcription through the deposition of H3K9me3 (Guang et al. 2010).

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Concomitant with a big wave in zygotic transcription, embryonic nuclei acquire methylated

H3K9me (Figure 3.1A), suggesting transcription elongation could be rate-limiting for

H3K9me2 deposition during embryogenesis. However, H3K9me2 levels were not affected after blocking transcription elongation (Figure 3.2), suggesting that it is not the rate-limiting component. This notion is supported by the fact that RNAi machinery is not required for the timing of H3K9me2. We note that while transcription elongation was undetectable under our experimental conditions, abortive transcription may still occur.

Thus, our results do not rule out transcription elongation or RNAi as a mechanism for targeting H3K9 methylation but rule them out as timers.

The nature of the relationship between zygotic transcription and chromatin regulation has been uncertain. One idea is that that chromatin regulation precedes and initiates zygotic transcription (Østrup, Andersen, and Collas 2013; Meier et al. 2018).

Conversely, zygotic transcription may help shape chromatin structure, a model tested by several studies: A study that analyzed the spatial genome organization in fly embryos revealed that early expressed genes correlate with the boundaries of Topologically

Associating Domains (TADs) (Hug et al. 2017). However, emergence of these structures are not dependent on transcription (Hug et al. 2017). Another study in Xenopus embryos mapped the distribution of histone modifications across the genome, and found that

H3K4me3 and H3K27me3 domains were established independently of zygotic transcription (Hontelez et al. 2015), similar to our results.

Classical models of time-keeping and H3K9 methylation

Classical models of embryonic time-keeping include i) an absolute clock that depends on the initiation of a signaling cascade and regulates the cell cycle (Howe and Newport 1996;

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Treen et al. 2018), ii) the increasing nuclear-to-cytoplasmic ratio (Newport and Kirschner

1982a; Pritchard and Schubiger 1996) or iii) the increasing DNA content in the embryo that titrates a maternal repressor (Newport and Kirschner 1982b; Dekens et al. 2003).

These models have mostly been studied in the context of zygotic genome activation. One study analyzed local changes in chromatin organization during fly embryogenesis and found that promoter accessibility is controlled by the nuclear-to-cytoplasmic ratio (Blythe and Wieschaus 2016). However, large-scale changes in chromatin organization during development such as heterochromatin formation had not been studied through the lens of these models.

In this study, we applied these models to large-scale chromatin organization. H3K9 methylation emerges at gastrulation and initiates heterochromatin formation in C. elegans embryos (Figure 3.1A, Chapter 2). To test how the timing of H3K9 methylation was dictated, we slowed down early embryonic cell divisions and uncoupled the number of cells from the amount of time spent after fertilization. Our results suggest that an absolute clock, rather than cell counting mechanisms, dictates the timing of H3K9 methylation.

Moreover, this absolute clock depends on the spatial regulation of MET-2 and its binding partners (Figure 3.4).

Spatial regulation of proteins provides a rapid means to restrict their activity. Prmt1 and SIRT1 each transition from the cytosol to the nucleus, or vice versa, to alter their activity upon differentiation (Hisahara et al. 2008; Ancelin et al. 2006). Similarly, MET-2 moves gradually into nuclei in the pre-gastrula embryo, but is released into the cytosol during mitosis (Chapter 2). Slowing down the cell cycle in div-1 embryos leads to precocious accumulation of MET-2 in the nucleus (Figure 3.4). Early embryonic cells

109 divide rapidly, with a 40-minute cell cycle, that likely interferes with the accumulation of

MET-2 in nuclei. Consistent with this idea, cells that divide slowly (the E cells) accumulate more nuclear MET-2 and more H3K9me2 than more rapidly dividing cells (data not shown).

Distinct roles for di vs. tri methylated H3K9 in cell fate potential

The differential effects of met-2 and set-25 on mono, di and tri methylated H3K9 provided a means to distinguish the roles of these histone marks. Developmental potential was inversely correlated with H3K9me2 (Figure 3.6): met-2, wild-type and set-25 embryos had low, average and high levels of H3K9me2, respectively. met-2 mutants were able to alter their developmental fate robustly, wild-type embryos less so, and set-25 mutants least of all. Double mutants between met-2 and set-25 lacked H3K9me2, and as predicted, they extended plasticity like met-2 single mutants. At later stages, met-2 mutants were resistant to the CFCA, suggesting that H3K9me2 is important for the timely loss of plasticity but not absolutely required. We note that our results do not rule out a role for

H3K9me1, but that H3K9me1 levels did not correlate with plasticity as closely as

H3K9me2 levels.

3.5 Materials and Methods

Strains. Strains were maintained at 20oC according to (Brenner 1974), unless stated otherwise*.

N2 (wild-type Bristol) *EU548 div-1(or148ts) III. (Encalada et al. 2000) WM161 prg-1(tm872) I. (Maintained at 15oC, experiments done at 20oC) YY186 nrde-2(gg91) II. (Guang et al. 2010) YY158 nrde-3(gg66) X. (Guang et al. 2008)

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WM126 sago-2(tm0894) ppw-1(tm0914) I; C06A1.4(tm0887), F58G1.1(tm1019) II; M03D4.6(tm1144) IV; sago-1(tm1195) V. (Yigit et al. 2006) *KW1975 taf-6.2(ax514); unc-17(e113) IV. SM2440 jmjd-1.2 (tm3713) IV. (Kleine-Kohlbrecher et al. 2010) EL597 omIs 1 [Cb-unc-119 (+) met-2::gfp II]. SM2575 lin-65::3xflag I. SM2333 pxSi01 (zen-4::gfp, unc-119+) II; unc-119(ed3) III. KM167 HS::hlh-1, (Fukushige and Krause 2005). SM1623 HS::hlh-1; met-2 (ok2307) III. JAC500 his-72(csb43[his-72::mCherry]) III, provided by John Calarco.

Antibody staining: Antibody staining was performed as described previously (Mutlu &

Mango, submitted) The following antibodies were used for immunostaining by 5min 2% paraformaldehyde (PFA), 3 min methanol (for all except PHA-4, which was fixed with

10min 2% PFA, 3min methanol). H5 Pol II staining followed a completely different protocol described in (L. Kaltenbach et al. 2000).

H3K9me2 (1:200) Abcam ab1220, MABI0307 Kimura 6D11 Histone H3 (1:500) Abcam ab1791 Pan-histone (1:500) Chemicon/Millipore MAB052 FLAG M2 (1:100) Sigma F1804 MET-2 (1:500) Raised against the first 17 amino acids of MET-2 and affinity purified, a gift from Eleanor Maine. ARLE-14 (1:500) Generated by our lab as described in (Mutlu & Mango, submitted). H5 Polymerase II (1:100) Covance MMS129-R Paramyosin (1:50) Developmental Studies Hybridoma Bank 5-23 PHA-4 N-terminus (1:1000) (L. S. Kaltenbach, Updike, and Mango 2005)

Quantitation of histone modifications and nuclear proteins. Analysis was done as described in Chapter 2. Briefly, embryos were imaged with a ZEISS LSM700 or LSM880

Confocal Microscope and analyzed by Volocity Software. Signal intensity of marks were calculated for each nucleus and average values for nuclei at designated embryonic stages plotted.

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Pouring RNAi plates. RNAi clones from the Ahringer library were used unless stated otherwise. First, identity of clones was confirmed by sequencing. To pour plates, bacteria were grown in 5ml LB with 5μl Carbenicillin (100mg/ml) for 6-8 hours at 37oC and pelleted at 4000rpm for 10 minutes. The bacterial pellet was resuspended in 400μl 0.5M IPTG,

30μl 100mg/ml Carb and 70μl Nuclease-free water. 5ml NGS plates were seeded with

100μl of resuspended bacterial solution and kept at room temperature for 2 days before use.

Transcription blocking. KW1975 taf-6.2(ax514) was maintained at 15oC. For experiments, KW1975 L4s were fed bacteria containing an ama-1 RNAi vector. In parallel,

SM2233 L4s were fed with bacteria containing empty vector. Both strains were grown at

15oC for 50-60 hours. Adult worms were dissected at 26oC and 1-4 cell embryos were transferred onto the same poly-L-lysine slide. Embryos were aged for 1 hour at 26oC in a humidity chamber and stained for H3K9me2. For H5 Polymerase II stains, wild-type

JAC500 worms fed with empty vector were used as an on-slide control instead of

SM2233. div-1 experiments. div-1(or148ts) was maintained at 15oC and shifted to 26oC for experiments. 2-cell stage wild-type zen-4::gfp (SM2233) and div-1 (EU548) embryos were picked and aged for 1 hour on the same poly-L-lysine slide in a humidity chamber at 26oC.

H3K9me2 staining after embryonic temperature shifts compared 25-30 cell wild-type embryos to 10-15 cell div-1 embryos.

Temperature shifts to 26oC that started with L4 animals instead of embryos produced identical results in terms of H3K9me2 levels at given embryonic stages. In L4-shift experiments, mixed stage SM2233 and EU548 embryos were dissected from gravid

112 adults and stained for H3K9me2 on the same slide. Cells that contained the same number of cells were compared using L4-shifts.

For atl-1 rescue experiments, N2 L4s were fed with atl-1 RNAi bacteria. div-1(or148ts)

L4s were fed with either empty vector or atl-1 RNAi bacteria overnight at 26oC. SM2233

L4s were fed with bacteria containing an empty vector at 26oC and included as an on- slide control on all experiments. Embryos were dissected from gravid adults and stained for H3K9me2.

As a control, bacteria containing the div-1 RNAi vector was fed to N2 and bacteria containing empty vector to SM2233 L4 animals overnight at 25oC. Embryos were dissected out of gravid adults and stained for H3K9me2 on the same slide (data not shown). These experiments gave identical results to div-1(or148ts) experiments.

CFCA. Cell Fate Challenge Assay was conducted similarly to (Kiefer et. al, 2007). In brief, two-cell embryos were collected from wild-type, set-25 or met-2 mothers carrying an integrated HS::hlh-1 array (Fukushige and Krause 2005). Embryos were incubated at

20oC for 3 hours until they reached the 100-cell stage, determined by DAPI staining and cell counts. Heat shock was administered at 33oC for 30 minutes on Poly-L-Lysine slides in a humidity chamber and embryos were incubated at 20oC for 20 hours. Terminally differentiated embryos were stained for paramyosin (muscle, (Fukushige and Krause

2005)) and PHA-4 (foregut, (Horner et al. 1998)). Embryos were imaged using the Zeiss

LSM 700 confocal microscope. RNA expression analysis for hlh-1 was done as described previously (Chapter 2).

GO Term Analysis. H3K9me1 (GSE49744), H3K9me2 (GSE49736) and H3K9me3

(GSE49732) methylated regions were defined by a MACS2 broad peak call (Zhang et al.

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2008) and the center of peaks were assigned to genes to curate a list. DAVID

(https://david.ncifcrf.gov/) was used for GO Term analysis.

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CHAPTER 4

CONCLUSION

4.1 Overview

In this chapter, I will first summarize my findings and discuss the novelty and significance of my results in the context of the field of chromatin organization. Then I will propose three follow-up projects based on the work described in this thesis and unpublished preliminary results: i) Characterizing LIN-65 and its upstream regulators, ii) Dissecting the role of the novel protein ARLE-14 in worms and humans, and iii) Understanding the role of germ granule localized MET-2.

4.2 Summary and Discussion

Chromatin re-organization is crucial for a cell to adapt to changing conditions and coordinate its transcriptional programs. Many developmental systems such as ESCs and animal embryos segregate their genome into euchromatin and heterochromatin domains during differentiation (Niwa 2007; Fadloun, Eid, and Torres-Padilla 2013; Politz, Scalzo, and Groudine 2013). While the complexes that re-organize chromatin are well studied, little is known about the temporal regulation of this process. This dissertation explored the molecular mechanisms behind the onset of heterochromatin formation, using C. elegans embryogenesis as a model.

Chapter 1 reviewed different methods that can be used to study chromatin organization, the changes they detected during development in different systems and highlighted C. elegans as a model organism to study large-scale chromatin organization in vivo during embryogenesis. During embryogenesis, chromatin accessibility is regulated locally at the level of individual nucleosomes (Teif et al. 2012; West et al. 2014), and globally at the level of chromatin domains: Early embryonic cells have large nuclei with a

123 uniform distribution of euchromatin (Niwa 2007). As cells mature, nuclei shrink, chromatin compacts, the genome is broadly segregated into euchromatin and heterochromatin

(Politz, Scalzo, and Groudine 2013), and methylated H3K9 domains emerge (Lepikhov and Walter 2004; Yeo et al. 2005; Wen, Wu, and Shinkai 2009; Yuan and O’Farrell 2016).

Heterochromatin is important for genome stability (McMurchy et al. 2017), regulation of gene expression and cell-fate restriction (Fadloun, Eid, and Torres-Padilla 2013).

C. elegans embryogenesis is a well-suited system to study heterochromatin formation because of the transparency, rapid development, ex utero survival and easy genetic manipulation of embryos. Moreover, it is challenging to study H3K9 methylation in vertebrates, since there are several partially redundant methyltransferases and the loss of each one has severe consequences (Allis and Jenuwein 2016). In worms, there are two main enzymes responsible for H3K9 methylation and embryos are viable in the absence of H3K9 methylation under ideal conditions (Towbin et al. 2012). This enables loss-of-function analysis to dissect the regulation of H3K9 methylation and provides insights into the process in mammalian cells.

Chapter 2 focused on the temporal regulation of the H3K9 methyltransferase MET-

2, an enzyme that is required for generating heterochromatin domains in C. elegans embryos. A proteomics approach identified two conserved binding partners that regulate

MET-2 and accumulation of H3K9 methylation during differentiation. One of these factors,

ARLE-14, was an unnamed worm protein (B0336.5) and the role of its human ortholog

ARL14EP in the nucleus is completely unknown (Paul et al. 2011). This is the first study that describes a role for ARLE-14 in H3K9 methylation in any organism.

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Poly Cystic Ovary Syndrome (PCOS) is one of most common disorders amongst women of reproductive age, with symptoms that significantly impact quality of life: menstrual dysfunction, androgen excess and an increased risk of obesity, diabetes and cancer (El Hayek et al. 2016). A genome-wide association study (GWAS) has implicated

ARL14EP in PCOS (Hayes et al. 2015), making my work potentially relevant to human disease. The 11p14.1 locus associated with PCOS spans an extensive 129.5 kb region beginning upstream of FSHB (follicle stimulating hormone beta subunit) and continues through ARL14EP (Hayes et al. 2015). Another study found differential gene expression and DNA methylation for ARL14EP in adipose tissue from women with PCOS compared to healthy women, but no changes in the FSHB expression or methylation (Kokosar et al.

2016), making ARL14EP the strongest candidate at the 11p14.1 locus. Moreover, lower levels of H3K9me2 have been observed in some studies of PCOS (Eini et al. 2017). My results suggest and intriguing link between ARL14EP and PCOS, potentially through regulation of H3K9 methylation.

I identified another conserved binding partner of MET-2: LIN-65, a worm protein that influences H3K9 methylation. Homologs of LIN-65 in non-nematode systems were elusive due to its disordered, acidic nature since its first molecular characterization in

2006 (Ceol et al. 2006; Tian et al. 2016). Through secondary structure predictions and functional analysis, I identified human ATF7IP and fly Windei as similar proteins that regulate localization of SETDB1/MET-2 (Koch et al. 2009; Timms et al. 2016).

Human ATF7IP is thought to both move SETDB1 into nuclei and shield nuclear

SETDB1 from degradation in the nucleus (Timms et al. 2016). The latter conclusion relies on the fact that nuclear SETDB1 is recovered in cells without ATF7IP after treatment with

125 proteasome inhibitors. The caveat is that proteasome inhibitors effect not only SETDB1, but degradation of every other protein in the cell. In worms, it is unlikely that LIN-65 shields

MET-2 from degradation in the nucleus, because i) total levels of MET-2 remain unchanged in lin-65 mutants, and ii) adding a strong NLS to MET-2 is sufficient to bypass the requirement for LIN-65 (Chapter 2). A more likely scenario is that LIN-65 affects the subcellular distribution of MET-2 by either blocking export of MET-2 or actively shuttling

MET-2.

LIN-65 influences both MET-2 accumulation in the nucleus and deposition of H3K9 methylation by MET-2, suggesting that it’s a critical factor for initiating heterochromatin formation in embryos. Intriguingly, LIN-65 is rate-limiting for not only H3K9me2, but also

H3K9me3, demonstrated by loss-of-function and half-dose LIN-65 experiments (Chapter

2). This is consistent with the step-wise model of H3K9 methylation, where MET-2 deposits H3K9me1/me2 and H3K9me2 is converted into H3K9me3 by SET-25 (Towbin et al. 2012).

Chapter 3 analyzed how embryonic cells keep time to initiate heterochromatin formation. I systematically tested whether any of the classical time-keeping mechanisms in developmental biology regulated chromatin re-organization during development. One classical model involves the presence of a repressor in excess in the early embryo and its titration during embryonic cell divisions, by the increasing ratio between nuclear and cytoplasmic volume (Newport and Kirschner 1982a; Pritchard and Schubiger 1996) or the increase in DNA content (Newport and Kirschner 1982b; Dekens et al. 2003).

Alternatively, the amount of time spent after fertilization could be a timer. During oocyte maturation, entry into mitosis triggers a cascade of events that eventually results in the

126 degradation of cyclins (Howe and Newport 1996; Treen et al. 2018) and slowing down of cell divisions (Kimelman, Kirschner, and Scherson 1987). Models of cell counting and absolute time are not mutually exclusive. For instance, in Drosophila embryos, most of zygotic transcription is activated based on absolute time after fertilization, whereas activation of a small class of genes relies on the nuclear-to-cytoplasmic ratio (Lu et al.

2009).

To distinguish between models of cell counting vs. absolute time, I slowed down early cell divisions (Encalada et al. 2000) and uncoupled number of cells and the amount of time after fertilization. My results suggest that the timing of H3K9 methylation is independent of cell counting mechanisms. Instead, it is dictated by an absolute clock which influences nuclear accumulation of MET-2 and its binding partners (Chapter 3).

The molecular nature of this absolute clock remains an unanswered question. One candidate was the Mitogen Activated Protein Kinase (MAPK) signaling cascade which becomes activated at fertilization (Miller et al. 2001). However, my preliminary results suggest that the deletion of the MAPK-kinase MEK-1 in C. elegans embryos has no effect on H3K9me2 (data not shown). It possible that similar to frog and ciona embryos (Howe and Newport 1996; Treen et al. 2018), a cyclin is gradually depleted in worm embryos at pre-gastrula stages and regulates accumulation of MET-2 and its binding partners in the nucleus, either by directly regulating these proteins or through the extension of interphase. The worm homolog of the regulated frog cyclin is cye-1 (cyclin E) (Fay and

Han 2000). cye-1 promotes the transition from G1 to S-phase in the germ line (Fox et al.

2011), however its role in embryonic cell divisions is less clear. A genome-wide single

127 cell map of mRNA abundance in pre-gastrula worm embryos suggests that cye-1 levels gradually decrease (Tintori et al. 2016), making it a good candidate.

In sum, this dissertation has made three contributions to scientific literature. First,

I found that regulated nuclear accumulation of a histone methyltransferase initiates heterochromatin formation during embryogenesis. Second, I have identified and characterized two conserved binding partners of this methyltransferase, with potential relevance to human disease. Third, I have dissected the molecular determinants that dictate the timing chromatin organization in embryos.

4.3 Future Directions i) Characterizing LIN-65 and its upstream regulators

• LIN-65: a disordered protein that organizes the nucleus

Membrane-less organelles are dynamic structures with liquid-like physical properties such as nucleoli in C. elegans. These properties arise from phase separation of their molecular components (Brangwynne et al. 2009; Brangwynne, Mitchison, and Hyman

2011). It is thought that intrinsically disordered protein regions (IDRs) that are conformationally heterogeneous and dynamic may mediate phase separation (Elbaum-

Garfinkle et al. 2015; Nott et al. 2015).

An emerging view is that phase separation also drives formation of heterochromatin compartments (Strom et al. 2017). I found that LIN-65, a protein that consists predominantly of disordered sequences, forms concentrated hubs in the nucleus together with MET-2. Without LIN-65, dilute amounts of MET-2 become nuclear, but hubs disappear (Appendix Figure 2). It is tempting to speculate that LIN-65 may be involved in

128 building the heterochromatin compartment by a phase-separation mechanism. SETDB1 is likewise found in concentrated hubs within cells (Cho et al. 2012), raising the speculation that the function of human ATF7IP or fly Windei is to build SETDB subdomains within nuclei.

Further work is needed to test this hypothesis in worms. The first step would be to assay if MET-2 or LIN-65 hubs behave as liquid droplets. There are several approaches to test whether proteins behave as liquid droplets in embryonic nuclei. First, live imaging of fluorescently labeled MET-2 or LIN-65 over 1 second time intervals would show whether these hubs fuse together or separate from each other, like phase-separated droplets would (Saha et al. 2016). Second, phase-separation is temperature dependent, where colder temperatures induce condensation (Nott et al. 2015). If MET-2 or LIN-65 hubs behave as liquid droplets, one would expect changes in temperature to alter their formation. Live imaging of these proteins at different temperatures would determine whether they behave as liquid droplets. Third, formation of liquid droplets are concentration dependent (Lin et al. 2015; Smith et al. 2016). In vitro phase-separation assays that use purified protein provide insights into the nature of these hubs in vivo.

Purified MET-2 or LIN-65 protein can be seeded at different concentrations and imaged to see if formation of hubs is concentration dependent. Concentration of MET-2 and LIN-

65 in embryonic extracts have been measured to be 60nM and 140 nM respectively (Saha et al. 2016), which could be used to determine a physiologically relevant range for in vitro experiments.

The second step would be to characterize the role of LIN-65 in the formation of MET-

2 hubs, and vice versa. My preliminary results are encouraging yet inconclusive. Forcing

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MET-2 into the nucleus in the absence of LIN-65 did not rescue H3K9 methylation

(Chapter 2), suggesting that LIN-65 is important for regulating nuclear MET-2 as well.

However, more careful analysis is required to understand how MET-2 hub formation is affected in the absence of LIN-65. Live imaging combined with in vitro experiments to analyze the behavior of MET-2 hubs in the presence and absence of LIN-65 would be informative.

• Structure-function analysis for LIN-65

Independent from a potential role in phase-separation, there’s still a lot to be discovered about LIN-65. It would be useful to do a structure-function analysis with LIN-

65 to determine minimal regions that are required for MET-2 binding and H3K9me2. LIN-

65 is mostly disordered but has two predicted domains based on my analysis. Intriguingly,

LIN-65 has three isoforms: one full length, and two isoforms that contain each domain separately. It would be interesting to see whether the function of different domains can be uncoupled in the cell through these different isoforms during development. It would also be interesting to characterize the role of these different domains in phase-separation, if applicable.

• Upstream regulators of LIN-65

Another remaining question is what acts upstream of LIN-65. My results suggest that

LIN-65 is rate-limiting for both H3K9me2 and H3K9me3, and critical for MET-2 localization and activity (Chapter 2). Early div-1 mutant embryos with precocious H3K9me2 also have more LIN-65 in the nucleus compared to wild-type (Chapter 3). While I’ve shown that an absolute clock is critical for establishing H3K9me (Chapter 3), it’s unclear what regulates

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LIN-65 during embryogenesis on a molecular level. It could be that LIN-65 is translationally regulated, post-translationally modified, or guided by a cell cycle protein.

While analyzing the post-translational modifications on MET-2 and LIN-65 protein in early vs. late embryos would be very interesting, it is technically challenging for two reasons: i) Protocols for getting perfectly synchronized embryo populations typically yield

300 thousand embryos, and ii) most proteomics approaches require 5-10 million embryos to get enough material. Instead, looking for binding partners of LIN-65 by co-IP and mass spectrometry may provide some insights into this process. iii) Dissecting the role of the novel protein ARLE-14 in worms and humans

• Upstream regulators of ARLE-14

ARLE-14 is a novel worm protein that is required for H3K9 di-methylation. My results show that ARLE-14 promotes association of MET-2 with chromatin (Chapter 2), but it’s unclear whether other pathways feed into the ARLE-14/MET-2 interaction to regulate chromatin organization. ARLE-14 is orthologous to vertebrate ARL14EP (ADP- ribosylation factor(ARF)-like Effector Protein). ARF family proteins undergo a cycle of

GTP binding and hydrolysis, which regulates their interactions with Effector Proteins (EP) that initiate downstream signaling cascades (Donaldson and Jackson 2011). It is tempting to speculate that ARLE-14/ARL14EP is regulated by an upstream ARF-like (ARL) protein in worms. A co-IP and mass spectrometry approach could identify factors that regulate

ARLE-14 and provide insights into the elusive role of ARL family proteins in the nucleus

(Burd, Strochlic, and Setty 2004).

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• Conservation of ARLE-14 in humans and its role in disease

A nuclear function for human ARL14EP had not been described, but three lines of evidence suggest ARL14EP functions within nuclei. First, ARL14EP was found to interact with SETDB1 in large-scale screens in humans and flies (Giot et al. 2003; Guruharsha et al. 2011) and by co-immunoprecipitation (Chapter 2). Second, ARL14EP is detected in nuclei of human tissues (www.proteinatlas.org) and C. elegans embryos (Chapter 2).

Third, C. elegans arle-14 is required to produce normal levels of H3K9me2 (Chapter 2).

Both ARL14EP and H3K9 methylation have been implicated in PCOS but were considered independent aspects of the disease (Hayes et al. 2015; Kokosar et al. 2016;

Eini et al. 2017). My results suggest an intriguing link between ARL14EP and H3K9 methylation in PCOS. It would be interesting to study the relationship between ARL14EP and SETDB1 in human cell lines and PCOS patient samples. For starters, are the levels or distribution of SETDB1/H3K9 methylation different in human cell lines without

ARL14EP? Is ARL14EP expression or genomic SETDB1/H3K9me distribution altered in

PCOS patient samples compared to healthy tissues? iii) Understanding the role of germ granule localized MET-2.

Germ cells are passed on from one generation to the next and have the potential to proliferate indefinitely. Germ granules are large, membraneless ribonucleoprotein (RNP) organelles found in the cytosol of most animals and are thought to be the key determinants of germ cell identity (Chuma et al. 2009). Many different types of granules exist, including perinuclear granules, stress bodies and processing bodies (P-bodies) and

132 they all contain a mixture of RNA with shared and unique protein components (Voronina et al. 2011).

In C. elegans, many factors involved in RNA silencing localize to germ granules

(P-granules) and are thought to surveil mRNAs as they exit the nucleus (Updike and

Strome 2010; Phillips et al. 2012; Wang and Seydoux 2014). Adjacent to P-granules are mutator foci, named after the mutator proteins that localize to these granules and are required for siRNA amplification and transposon silencing (Zhang et al. 2011).

Intriguingly, I noticed that the histone methyltransferase MET-2 localizes to germ cell granules during mitosis (Figure 4.1), and I detected low amounts of germ granule components as MET-2 binding partners in my proteomics analysis (Appendix Table A.1-

2). It is unclear whether these are P-granules, mutator foci or another type of granule. Co- staining experiments with unique granule components such as PGL-1 or MUT-16 will reveal the type of granules MET-2 localizes to.

Figure 4.1. MET-2 localizes to germ cell granules during mitosis. 8-cell stage embryo showing the germ (P) cell during mitosis. Antibody staining against MET-2::GFP (green), DAPI (blue). White arrrow points to the granules. Scalebar 2μm.

MET-2 is absent from granules during interphase but appears during mitosis. This could be due to two reasons: First, MET-2 is released from the nucleus as cells divide, increasing the cytosolic pool of MET-2 (Chapter 2). Second, P-granules fuse with each

133 other and grow in size during mitosis (Gallo et al. 2010), which could promote stable associations with MET-2. Studying dynamics of MET-2 could provide insights into the mechanism of localization to germ granules.

The role of granule associated MET-2 is mysterious and could be germ-cell specific. It may be related to its mainstream role in silencing repetitive sequences in the adult germline (Zeller et al. 2016; McMurchy et al. 2017), and these granules may serve as scaffolds that assemble silencing complexes. In support of this idea, my preliminary results show that RNA for a tandem repeat silenced by MET-2 also localizes to granule- like structures in the embryonic P-cell (data not shown). It remains to be tested whether granule associated MET-2 eventually becomes nuclear. This could be done by utilizing photoswitchable tags to track intracellular MET-2 movements (Chudakov, Lukyanov, and

Lukyanov 2007). Alternatively, MET-2 may have a completely independent role and methylate non-histone substrates in these granules. The role of cytosolic MET-2 and its potential non-histone substrates is being studied as a separate project in our laboratory, in collaboration with Dr. Jessica Tanis.

4.4 Concluding Remarks

Chromatin re-organization is a critical process for cancer (Zink, Fischer, and

Nickerson 2004), numerous diseases (Hendrich and Bickmore 2001) and embryogenesis

(Politz, Scalzo, and Groudine 2013). My work has focused on dissecting the molecular cues that determine the onset of chromatin organization during embryogenesis and has revealed that a conserved histone methyltransferase and its binding partners regulate this transition. Future work that further characterizes the role of these factors may contribute

134 to emerging fields such as compartmentalization of the nucleus through phase- separation, identify new roles for well-studied proteins, and shed light on molecular processes involved in human disease.

4.5 References

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Brangwynne, Clifford P, Timothy J Mitchison, and Anthony A Hyman. 2011. “Active Liquid-like Behavior of Nucleoli Determines Their Size and Shape in Xenopus Laevis Oocytes.” Proceedings of the National Academy of Sciences of the United States of America 108 (11). National Academy of Sciences:4334–39. https://doi.org/10.1073/pnas.1017150108.

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Chudakov, Dmitriy M, Sergey Lukyanov, and Konstantin A Lukyanov. 2007. “Tracking Intracellular Protein Movements Using Photoswitchable Fluorescent Proteins PS- CFP2 and Dendra2.” Nature Protocols 2 (8). Nature Publishing Group:2024–32.

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Donaldson, Julie G., and Catherine L. Jackson. 2011. “ARF Family G Proteins and Their Regulators: Roles in Membrane Transport, Development and Disease.” Nature Reviews Molecular Cell Biology 12 (6). Nature Publishing Group:362–75. https://doi.org/10.1038/nrm3117.

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APPENDIX

Supplementary Figures for Chapter 2

Supplementary Figure S2.1 A. TEM single sections of whole embryos at designated stages. Each embryo is approximately 50μm long. B. Whole embryos stained with H3K9me1/2/3, HK27me3 or H4 pan-acetylation at different stages of development (Scale bar 2 μm). C. Antibody specificity for H3K9me2: representative single nuclei at designated stages showing H3K9me2 staining with two additional H3K9me2 antibodies: Upstate 07-441 and Kimura 6D11. D. Antibody specificity for H3K9me3: representative single nuclei at designated stages showing H3K9me staining in wild-type vs. set-25 mutant embryos, Histone Modification (HM, green), DAPI (blue). H3K9me/Histone levels normalized to wild-type.

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Supplementary Figure 2.2 A. TEM single sections of whole met-2 embryos. Note the electron dense droplets in the cytoplasm vs. the electron lucent nuclei. B. Cytosolic components in WT vs. met-2 mutants by TEM. Mitochondria (red circle), endoplasmic reticulum (blue box), lipid droplet (purple arrowhead). Note the similar

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Supplementary Figure 2.2 continued appearance of the cytosol vs. the different morphologies of the nuclei for wild-type vs met- 2. C. H3K9me2 staining in wild-type vs. his-72::mCherry embryos showing that the mCherry tag doesn’t interfere with H3K9me2. D. Whole embryos stained with an antibody against endogenous MET-2 (raised against the first 17 amino acids of MET-2 protein) and pan-histone. (-) represents met-2 mutants. Scale bar, 2 μm. E. Representative single nuclei showing Crispr reporter 3xFLAG::MET-2 during interphase (INT) and prophase (PRO, Scale bar, 2 μm). F. H3K9me2 levels in interphase (INT) and prophase (PRO) nuclei from the same embryo at the 15 cell stage.

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Supplementary Figure 2.3 A. Table showing spectral counts and peptide coverage after GFP IP-MudPIT MS for the following strains: met-2::gfp, wild-type (no GFP) and zen-4::gfp (GFP control). B. Silver stain showing the 100 kDa LIN-65 band after MET-2::GFP IP, identified by cutting out the 100kDa band and Mass Spec. C-D. Whole embryos showing LIN-65::3xFLAG (FLAG antibody; C) and endogenous ARLE-14 (D) at different stages of embryonic development with a H3 or pan-histone co- stain (Scale bar, 2 μm). Quantitation of nuclear and total protein during embryogenesis, normalized to histone. E. Left panel shows the staining pattern of transcription factor PHA-4::GFP (green) in whole embryos, with DNA (DAPI). The two panels on the right show PLA signal between GFP and pan-histone (red) with DNA (DAPI) in the following strains: pha-4::gfp or wild- type N2 (no GFP -PHA-4::GFP) (Scalebar 2 μm).

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Supplementary Figure 2.4

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Supplementary Figure 2.4 continued

Supplementary Figure 2.4 A. Whole embryos showing the distribution of MET-2::GFP in WT vs. arle-14 mutants (Scale bar 2 μm). Line scan analysis showing the mean MET-2::GFP intensity across embryonic cells in WT vs. arle-14 mutant embryos. Average of line scans across multiple nuclei are shown and error bars denote standard error of the mean. B. Whole embryos stained with antibodies against endogenous MET-2 protein in wild- type (green) vs. lin-65 (red) mutants, and co-stained for pan-histone. C. MET-2::GFP IP and Western blot with GFP and histone antibodies in wild-type vs. lin- 65 embryonic extracts on the same gel. The lane between wild-type vs. lin-65 is not shown. The graph shows pixel counts for the intensity of MET-2::GFP normalized to histone H3. D. MET-2::GFP levels in progeny of met-2(+/-) moms. Quantitation of MET-2::GFP signal intensity in the cytosol and the nucleus. E. H3K9me2 levels in progeny of met-2(+/-) and arle-14(+/-) moms and quantified. F. MET-2::GFP levels in WT vs. progeny of lin-65(+/-) heterozygous moms. Line scan quantitation showing mean accumulation of MET-2::GFP in wild-type (grey) or lin-65/+ (purple) embryos at the 21-50 cell stage. Average of line scans across multiple nuclei are shown and error bars denote standard error of the mean. G. Coiled-coil probability across LIN-65 amino acid sequence predicted by PCOILS (https://toolkit.tuebingen.mpg.de/#/tools/pcoils).

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Supplementary Figure 2.4 continued H. Model structure of LIN-65 C-terminus predicted by Phyre2 (http://www.sbg.bio.ic.ac.uk/phyre2/html/page.cgi?id=index). Full LIN-65 a.a. sequence was used as input for Phyre2. I. Domain architecture from HMMR (https://www.ebi.ac.uk/Tools/hmmer/) for human ATF7IP, fly Windei and worm LIN-65 showing disordered (purple), coiled-coil (pink), and beta-sandwich fold (violet) regions. J. Amino acid sequence alignment of worm ARLE-14 to the human ARL14EP domain using ClustalX2. Color scheme: Hydrophobic-blue, Positive charge-red, Negative charge- magenta, Polar-green, Cysteines-pink, Glycines-orange, Prolines-Yellow, Aromatic-cyan. K. Protein interaction map for human ARL14EP from String database (https://string- db.org/). Red box highlights interactions with human SETDB1/2. L. ARLE-14 antibody staining in wild-type vs met-2, lin-65 and arle-14 mutants. Line scan quantitation. Average of line scans across multiple nuclei are shown and error bars denote standard error of the mean.

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Table A.1 - List of MET-2::GFP interactors zen-4::gfp and N2 embryos that lack a GFP tag were used as specificity controls.

Spectral Counts Coverage Accession Gene MET-2 ZEN-4 N2 MET-2 R05D3.11 met-2 895 7 0 35.90% Y71G12B.9a lin-65 171 0 0 21.00% B0336.5b arle-14 8 0 0 20.60% B0336.5a arle-14 8 0 0 20.90% C05C8.4 gei-6 (gex interacting protein) 216 3 0 17.90% C18G1.4b pgl-3 (P-granule component) 8 0 0 5.10% R144.7a larp-1 (P-bodies) 5 0 0 3.30% mbf-1 (Helix-turn-helix H21P03.1 Transcription factor) 8 0 0 17.90% icd-2 (NACA, Y65B4BR.5a methyltransferase interactor) 18 0 0 24.60% C27B7.1a spr-2 (SET domain protein) 6 0 0 6.40% rad-23 (RAD23 protein ZK20.3 homolog2 like) 6 0 0 3.10% hmg-1.1 (High Mobility Group Y48B6A.14 protein) 4 0 0 9.50% snr-5 (small nuclear ZK652.1 ribonucleoprotein Sm F) 4 0 0 21.20% T24F1.1 raga-1 (GTP-binding protein) 3 0 0 4.20% ola-1 (Obg like ATPase, W08E3.3b pred.GTP binding) 10 0 0 17.60% K06C4.5 his-17 (histone H3) 291 0 0 6.60% F45E1.6 his-71 (histone H3) 291 0 0 6.60% ZK131.3 his-9 (histone H3) 291 0 0 6.60% ZK131.2 his-25 (histone H3) 291 0 0 6.60% ZK131.7 his-13 (histone H3) 291 0 0 6.60% F45F2.13 his-6 (histone H3) 291 0 0 6.60% F54E12.1 his-55 (histone H3) 291 0 0 6.60% K06C4.13 his-27 (histone H3) 291 0 0 6.60% K03A1.1 his-40 (histone H3) 291 0 0 6.60% Y49E10.6a his -72 (histone H3) 291 0 0 6.60% F07B7.5 his- 49 (histone H3) 291 0 0 6.60% F55G1.2 his-59 (histone H3) 291 0 0 6.60% F17E9.10 his-32 (histone H3) 291 0 0 6.60%

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Table A.1 continued

T10C6.13 his-2 (histone H3) 290 0 0 6.60% B0035.10 his-45 (histone H3) 290 0 0 6.60% F22B3.2 his-63 (histone H3) 290 0 0 6.60% F08G2.3 his-42 (histone H3) 290 0 0 6.60% rpl-7A (Large ribosomal Y24D9A.4c subunit) 67 0 0 14.30% C54C6.1 rpl-37 (60S ribosomal protein) 15 0 0 7.70% eef-1G (translation elongation F17C11.9c factor) 53 0 0 9.30% Y71A12B.1c rps-6 (40S ribosomal subunit) 3 0 0 12.50% Y71H2AM.2 tufm-1 (mitochondrial 3 translation factor) 11 0 0 3.40% sars-1 (seryl-tRNA C47E12.1 synthetase) 16 0 0 8.40% dlat-1 (pyruvate F23B12.5 dehydrogenase complex) 10 0 0 2.40% pdha-1 (pyruvate T05H10.6a dehydrogenase subunit) 10 0 0 5.80% pdha-1 (pyruvate T05H10.6b dehydrogenase subunit) 10 0 0 5.60% K10C2.4 fah-1 (fumarylacetoacetase) 7 0 0 6.20% gpi-1 (glucose-6-phosphate Y87G2A.8a isomerase) 4 0 0 1.80% F32B5.8 cpz-1 (cysteine proteinase) 4 0 0 7.80% acdh-3 (acyl-CoA K06A5.6 dehydrogenase) 3 0 0 4.80% sucl-1 (succinyl-CoA C05G5.4 synthetase) 4 0 0 3.70% qars-1 (glutaminyl (Q) tRNA Y41E3.4b synthetase) 8 0 0 3.60% acdh-12 (acyl coA E04F6.5b dehydrogenase) 7 0 0 8.80% ttx-7 (myo-inositol-1- F13G3.5a monophosphatase) 7 0 0 5.60% B0432.2 djr-1.1 (glyoxylase) 6 0 0 12.80% Y34D9A.6 glrx-10 (glutaredoxin) 6 0 0 12.40% Y49E10.2 glrx-5 (glutaredoxin) 3 0 0 19.70% T24A11.1a mtm-3 (lipid phosphatase) 5 0 0 5.00% W07E11.1a glutamate synthase 8 0 0 1.60% Y110A7A.6b phosphofructokinase 8 0 0 10.60% C32F10.8a alanine aminotransferase 9 0 0 7.30%

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Table A.1 continued

ttr-50 (Transthyretin-like F58B3.9 family) 9 0 0 10.30% ZC155.1 nex-1 (annexin) 9 0 0 10.20% csn-5 (COP9 signalosome B0547.1 complex) 5 0 0 2.70% aldo-2 (fructose-bisphosphate F01F1.12b aldolase) 5 0 0 10.40% dod-23 (downstream of daf- F49E12.2 16) 5 0 0 12.50% uev-1 (ubiquitin conjugating F39B2.2 enzyme) 4 0 0 14.40% tag-72 (Yeast ABD1 protein C25A1.3 like) 4 0 0 5.80% F57B10.11 bag-1 (co-chaperone) 3 0 0 12.90% F01F1.8b cct-6 (chaperonin) 3 0 0 4.20% unc-11 (clathrin adaptor C32E8.10b protein) 3 0 0 3.30% B0546.1 mai-2 (ATPase inhibitor) 3 0 0 14.70% clpf-1 (poly-adenylation F59A2.4a factor) 2 0 0 4.90% ndk-1 (nucleoside F25H2.5 diphosphate kinase) 196 17 39 26.80% nasp-2 (histone binding C50B6.2 protein) 179 21 42 26.80% F25B5.4a ubq-1 (ubiquitin C) 157 0 64 4.50% Table A.1. List of MET-2::GFP binding partners.

Table A.2 - List of 3xFLAG::MET-2 interactors pha-4::gfp::3xflag (SM1754) and N2 embryos that lack a FLAG tag were used as specificity controls.

Spectral Counts Coverage Accession Gene MET-2 N2 PHA-4 MET-2 R05D3.11 met-2 287 0 0 29.10% Y71G12B.9a lin-65 167 0 0 13.50% Y71G12B.9b lin-65 133 0 0 11.50% C05C8.4 gei-6 76 0 0 31.10%

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Table A.2 continued

lin-53 (chromatin assembly K07A1.12 factor) 13 0 0 10.80% D2030.6 prg-1 (piwi pathway) 8 0 0 10.70% nrde-3 (nuclear RNAi R04A9.2 pathway) 11 0 0 3.10% C18E3.7c ppw-1 (RNAi argonaute) 11 0 0 4.40% F48F7.1a alg-1 (miRNA pathway) 5 0 0 2.60% T07D3.7 alg-2 (miRNA pathway) 7 0 0 4.20% F55A12.1 wago-2 (RNAi argonaute) 4 0 0 4.90% F56A6.1a sago-2 (RNAi argonaute) 4 0 0 2.70% K12H4.8 dcr-1 (RNAi pathway) 4 0 0 2.80% F22D6.6 ekl-1 (RNAi pathway) 2 0 0 3.60% F32H2.3 spd-2 (spindle defective) 12 0 0 3.90% T05G5.3 cdk-1 (cell cycle kinase) 10 0 0 17.50% mat-1 (anaphase promoting Y110A7A.17a complex) 5 0 0 6.50% mat-2 (anaphase promoting W10C6.1 complex) 2 0 0 1.10% F29F11.6a gsp-1 (protein phosphatase) 9 0 0 8.20% B0205.7 kin-3 (casein kinase) 5 0 0 11.10% K07C11.2 air-1 (protein kinase) 3 0 0 3.70% F35G12.3b sel-5 (serine threonine kinase) 3 0 0 3.60% K08A8.1a mek-1 (MAP kinase kinase) 2 0 0 7.80% par-1 (serine threonine H39E23.1a kinase) 2 0 0 3.40% egl-4 (cyclic-GMP-dependent F55A8.2a protein kinase) 2 0 0 3.10% B0261.2a let-363 (kinase) 2 0 0 0.90% B0041.2b ain-2 37 0 0 4.30% ZK418.9a possible RNA binding protein 22 0 0 8.40% K02B9.1 meg-1 (P-granule component) 3 0 0 3.50% deps-1 (P-granule Y65B4BL.2 component) 4 0 0 7.40% ZK858.1 gld-4 (P-granule component) 3 0 0 3.70% AH6.5 mex-6 (germline protein) 5 0 0 7.90% xpo-1 (exportin, nuclear ZK742.1b transport factor) 3 0 0 31.60% F32H2.4 thoc-3 (Tho-Trex xomplex) 5 0 0 6.00% C54G10.2a rfc-1 (replication factor C) 3 0 0 3.90% F33H2.5 DNA polymerase family B 3 0 0 1.00% F31E3.3 rfc-4 (Replication factor C) 2 0 0 6.90%

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Table A.2 continued

F58F6.4 rfc-2 (DNA replication factor) 2 0 0 7.80% C29A12.3a lig-1 (DNA ligase) 4 0 0 6.70% rpb-3 (DNA directed RNA C36B1.3 polymerase II) 3 0 0 6.70% cri-3 (Splicing factor- F59A2.3 associated) 4 0 0 21.20% rsp-2 (pre-mRNA splicing W02B12.2 factor like) 4 0 0 5.70% msh-2 (DNA mismatch repair H26D21.2 protein) 7 0 0 4.80% R07E5.8 cku-80 (DNA repair - NHEJ) 3 0 0 4.80% Y38C9A.2 cgp-1 (GTP-binding protein) 2 0 0 5.70% T24F1.1 raga-1 (GTP-binding protein) 2 0 0 7.10% F45E1.6 his-71 (histone H3) 3 0 0 8.10% T10C6.13 his-2 (histone H3) 3 0 0 8.10% ZK131.3 his-9 (histone H3) 3 0 0 8.10% ZK131.2 his-25 (histone H3) 3 0 0 8.10% ZK131.7 his-13 (histone H3) 3 0 0 8.10% K06C4.5 his-17 (histone H3) 3 0 0 8.10% F45F2.13 his-6 (histone H3) 3 0 0 8.10% F54E12.1 his-55 (histone H3) 3 0 0 8.10% K06C4.13 his-27 (histone H3) 3 0 0 8.10% K03A1.1 his-40 (histone H3) 3 0 0 8.10% Y49E10.6a his-72 (histone H3) 3 0 0 8.10% W09H1.2 his-73 (histone H3) 3 0 0 8.40% W05B10.1 his-74 (histone H3) 3 0 0 8.10% F08G2.3 his-42 (histone H3) 3 0 0 8.10% F17E9.10 his-32 (histone H3) 3 0 0 8.10% B0035.10 his-45 (histone H3) 3 0 0 8.10% F22B3.2 his-63 (histone H3) 3 0 0 8.10% F07B7.5 his-49 3 0 0 8.10% F22F1.1 hil-3 (histone H1) 4 0 0 11.10% T05C3.5 dnj-19 (DNAJ-like protein) 10 0 0 7.50% T24H10.3 dnj-23 (DNAJ protein) 4 0 0 13.60% chd-3 (helicase-DNA-binding T14G8.1 like protein) 9 0 0 2.60% snr-4 (small nuclear C52E4.3 ribonucleoprotein D2 like) 6 0 0 13.60% Y71F9AL.13b rpl-1 (large ribosomal subunit) 18 0 0 8.40% F28C6.7a rpl-26 (ribosomal protein) 4 0 0 14.10%

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Table A.2 continued

W09C5.6b rpl-31 (ribosomal proten) 3 0 0 17.10% ife-1 (translation initiation F53A2.6b factor) 8 0 0 6.10% ife-3 (translation initiation B0348.6a factor) 4 0 0 16.90% D2085.3 translation initiation factor 3 0 0 2.90% F57B9.3 Eukaryotic initiation factor 4A 7 0 0 5.80% H19N07.1a erfa-3 (elongation factor) 5 0 0 7.00% cars-1 (aminoacyl-tRNA Y23H5A.7a synthetase) 4 0 0 3.90% fars-3 (phenylalanyl-tRNA F22B5.9 synthetase) 4 0 0 3.00% ccr-4 (alcohol dehydrogenase ZC518.3a transcription effector like) 17 0 0 7.80% idha-1 (isocitrate F43G9.1 dehydrogenase) 15 0 0 29.30% gpdh-2 (Glycerol-3-phosphate K11H3.1a dehydrogenase) 12 0 0 17.00% fkb-2 (peptidyl-prolyl cis-trans Y18D10A.19 isomerase) 9 0 0 23.10% Y110A7A.4 thymidylate synthase 6 0 0 8.00% gsr-1 (pyridine nucleotide- C46F11.2a disulphide oxidoreductase) 6 0 0 13.50% ZK669.4 lipoamide acyltransferase 6 0 0 8.70% pck-2 (phosphoenolpyruvate R11A5.4a carboxykinase) 5 0 0 6.30% Y38A8.2 pbs-3 (Peptidase) 5 0 0 10.80% F54B3.3 atad-3 (ATPase) 5 0 0 9.20% F58F12.1 ATP synthase 6 0 0 20.90% W07E11.1a glutamate synthase 5 0 0 1.00% K11D9.2a sca-1 (E1-E2 ATPase) 5 0 0 4.30% mccc-1 (carbomoyl-phosphate F32B6.2 carboxylase) 4 0 0 3.70% ubiquitin carboxyl-terminal T05H10.1 hydrolase 5 0 0 3.50% Y39E4A.3a Transketolase 5 0 0 11.40% F57B10.3a phosphoglycerate mutase 3 0 0 4.80% RNA adenosine deaminase D2005.1 like 3 0 0 12.40% cyn-3 (Peptidyl-prolyl cis-trans Y75B12B.5 isomerase) 3 0 0 17.90%

156

Table A.2 continued

pdha-1 (pyruvate T05H10.6a dehydrogenase complex) 3 0 0 5.50% pars-1 (Prolyl-tRNA T20H4.3a synthetase) 3 0 0 3.40% F32H2.5 fasn-1 (fatty acid synthase) 3 0 0 0.90% pck-2 (phosphoenolpyruvate R11A5.4c carboxykinase) 3 0 0 5.30% kat-1 (acetoacetyl-C0A T02G5.8 thiolase) 3 0 0 7.10% cpt-2 (carnitine R07H5.2a palmitoyltransferase II) 3 0 0 3.30% 3-hydroxyacyl-CoA B0272.3 dehydrogenase 3 0 0 10.40% cas-2 (adenylyl cyclase- C18E3.6 associated protein) 3 0 0 5.50% Y54G11A.6 ctl-1 (catalase) 2 0 0 8.70% Y54G11A.13 ctl-3 (catalase) 2 0 0 8.40% C02G6.1 peptidase 2 0 0 1.90% pyrroline-5-carboxylate M153.1 reductase 2 0 0 5.70% dhs-6 (Alcohol C17G10.8 dehydrogenase) 2 0 0 5.50% F23B12.8a bmk-1 (kinesin-like protein) 8 0 0 4.80% T12E12.4a drp-1 (dynamin-like protein) 5 0 0 3.30% R05D3.7 unc-116 (Kinesin heavy chain) 4 0 0 4.80% hum-9 (Myosin head motor Y11D7A.14 domain) 3 0 0 1.20% C41G7.2 klp-16 (kinesin) 2 0 0 3.90% T22C1.6 myosin 2 0 0 5.10% ZK520.4a cul-2 (cullin family) 4 0 0 4.00% CD4.6 pas-6 (protease) 4 0 0 11.90% pas-2 (proteosome D1054.2 component) 2 0 0 12.10% T12F5.3 glh-4 (helicase) 3 0 0 2.90% T04D1.4 chd-7 (helicase) 3 0 0 1.50% rnp-3 (U1 small nuclear K08D10.3 ribonucleoprotein A) 2 0 0 9.20% lis-1 (WD domain, G-beta T03F6.5 repeats) 3 0 0 10.40% skp-1 (Drosophila puff specific T27F2.1 protein BX42 like status) 3 0 0 4.90% C29E4.8 let-754 (Adenylate kinase) 3 0 0 9.60%

157

Table A.2 continued

lex-1 (TAT-binding homolog F11A10.1a like) 2 0 0 1.30% Clathrin adaptor complex F59E10.3 small chain 2 0 0 8.20% Y47G6A.20b rnp-6 (RNA-binding protein) 2 0 0 3.30% C03D6.4 npp-14 (nucleoporin) 2 0 0 1.90% Forkhead-associated (FHA) C01G6.5 domain 3 0 0 2.60% ornithine aminotransferse C16A3.10a precursor 3 0 0 7.10% ape-1 (P53-binding protein F46F3.4a like) 3 0 0 4.00% gpb-1 (guanine nucleotide- F13D12.7a binding protein) 3 0 0 5.90% Y37D8A.1 arx-5 9 0 0 10.40% Y22F5A.4 lys-1 13 0 0 16.40% Y110A7A.16 elpc-1 9 0 0 1.70% R07G3.5 pgam-5 8 0 0 6.30% C33H5.4a klp-10 7 0 0 7.10% F58B3.5c mars-1 7 0 0 12.60% K08E7.2 hsb-1 7 0 0 30.00% Y39G10AR.1 0 epg-2 6 0 0 7.50% W10G11.20 dnc-3 6 0 0 33.90% K07D4.3 rpn-11 6 0 0 6.70% T24A11.1a mtm-3 6 0 0 3.70% Y56A3A.1a ntl-3 6 0 0 4.00% F33D11.11 vpr-1 6 0 0 18.00% F09G2.9 attf-2 6 0 0 6.70% Y54E10BR.6 rpb-7 6 0 0 19.80% C04C11.2 arrd-25 6 0 0 6.00% F26F4.1 cee-1 6 0 0 9.10% Y48B6A.12 men-1 5 0 0 4.70% B0350.2g unc-44 5 0 0 4.70% Y110A2AL.13 pinn-1 5 0 0 19.30% Y14H12B.1a zinc-finger protein 5 0 0 12.00% W01B6.9 ndc-80 5 0 0 9.30% C02E11.1a nra-4 5 0 0 1.80% Y55F3AM.15 csn-4 4 0 0 6.60% Y75B7AL.4a rga-4 4 0 0 4.50%

158

Table A.2 continued

K11D9.2c sca-1 4 0 0 3.00% Y61A9LA.8 sut-2 4 0 0 3.50% ZC518.2 sec-24.2 (Yeast YIK9 like) 4 0 0 1.60% R07H5.8 adenosine kinase 4 0 0 5.60% C25B8.3c cpr-6 4 0 0 6.50% Y56A3A.32a wah-1 4 0 0 4.40% W06A7.3d ret-1 4 0 0 6.30% C36E6.3 mlc-1 3 0 0 17.60% C36E6.5 mlc-2 3 0 0 17.60% F37D6.2a Zinc finger, C2H2 type 3 0 0 7.00% Y63D3A.5 tfg-1 3 0 0 4.30% K12D12.2 npp-3 3 0 0 1.80% B0513.1a lin-66 3 0 0 5.40% Y75B7AL.4b rga-4 3 0 0 14.00% Y56A3A.1b ntl-3 3 0 0 9.60% C14A4.14 mrps-22 3 0 0 5.30% H04J21.3a gip-1 3 0 0 3.40% Y47D3A.26c smc-3 3 0 0 7.00% C07G1.5 hgrs-1 3 0 0 2.90% VW02B12L.3 ebp-2 3 0 0 16.70% M03C11.4 hat-1 3 0 0 7.30% F19B6.2a ufd-1 3 0 0 5.80% R10E12.1a alx-1 3 0 0 4.80% C35E7.1a vet-2 3 0 0 3.10% F47G6.4 spe-15 3 0 0 2.50% Y108G3AL.1 cul-3 3 0 0 3.00% Y37A1B.1a lst-3 3 0 0 2.10% C47D12.1a trr-1 3 0 0 0.60% C24G6.3 mms-19 3 0 0 3.10% Y45G12B.1c nuo-5 3 0 0 4.10% ZK1128.8b vps-29 3 0 0 15.70% C29E4.2 kle-2 3 0 0 1.90% F21H11.2a sax-2 3 0 0 0.90% Y67H2A.6a csn-6 3 0 0 5.40% Y71F9AM.5b nxt-1 3 0 0 12.20% H14A12.2b fum-1 3 0 0 7.60% T16G12.5 ekl-6 2 0 0 2.50% F53A2.4 nud-1 2 0 0 10.00% F18E2.3 scc-3 2 0 0 2.00%

159

Table A.2 continued

K07H8.10 RNA-binding protein 2 0 0 3.30% C32E8.11 ubr-1 2 0 0 1.40% C25G4.5 dpy-26 2 0 0 2.10% W06B4.3 vps-18 2 0 0 2.70% C52E12.1 zinc-finger containing protein 2 0 0 3.50% F13G3.4 dylt-1 2 0 0 15.10% F26F4.10c rars-1 2 0 0 19.40% C43E11.1 acin-1 2 0 0 4.10% F55C5.8 signal recognition particle 2 0 0 3.90% F08B4.5 pole-2 2 0 0 5.80% F56G4.3 pes-2.2 2 0 0 6.60% F56G4.2 pes-2.1 2 0 0 6.60% K10D2.3 cid-1 2 0 0 1.30% C45G3.5 gip-2 2 0 0 4.20% F25B5.2 nop-1 2 0 0 2.80% Y94H6A.9a ubxn-1 2 0 0 13.00% M176.3 chch-3 2 0 0 17.20% R09B3.2 RNA recognition motif 2 0 0 25.30% Table A.2. List of 3xFLAG::MET-2 binding partners.

Experimental differences for MET-2::GFP and 3xFLAG::MET-2 IP

To find binding partners for MET-2, I used two reagents: MET-2::GFP and 3xFLAG::MET-

2. The generation of these strains and the details of the IP protocol are described in the

Materials and Methods Section of Chapter 2. However, I’d like to point out the differences between the two experiments as the list of interactors showed some variation: i) The position and size of the tag: The GFP tag is bulky (~30kDa) and was at the C- terminus of MET-2, where the SET domain is located. Note that MET-2::GFP resulted only in a partial rescue of H3K9me2 levels, but the timing was intact. The 3xFLAG tag is much smaller and was inserted at the N-terminus of MET-2 and restored both H3K9me2

160 levels and timing fully. It is possible that the position and size of the tag influenced binding interactions with MET-2. ii) Staging of embryo samples: Mixed stage embryos were used for both experiments.

The met-2::gfp sample was slightly younger than the 3xflag::met-2 sample, which could account for the differences. The breakdown of embryonic stages was as follows:

• met-2::gfp – 1-10 cell (24%); 11-20 cell (21%), 21-50 cell (20%), 51-80 cell (17%),

81-100 cell (15%).

• 3xflag::met-2 – 1-20 cell (3%), 21-50 cell (14%), 51-80 cell (25%), 81-100 cell

(50%), 101-200 cell (8%). iii) Beads and elution conditions: To accommodate different tags, different magnetic beads were used and the elution conditions were altered accordingly. For MET-2::GFP,

GFP-TRAP magnetic agarose beads and a glycine elution was used. For 3xFLAG::MET-

2, the FLAG M2 antibody was coupled to magnetic Protein A/G beads and a 3xFLAG peptide elution was used. The rest of the protocol was the same. See Methods of Chapter

2 for details.

Conservation of MET-2 in other organisms

Human SETDB1 (Basavapathruni et al. 2016) and G9a (Rice et al. 2003; Tachibana et al. 2002) are two histone methyltransferases that can catalyze mono- and di-methylation, similar to worm MET-2 (Towbin et al. 2012). To determine which is more likely to be the ortholog of MET-2, I looked at the conservation of amino acid residues in the SET domain across different species. By sequence, MET-2 is most similar to vertebrate SETDB1

(Appendix Figure 1A, (Poulin et al. 2005; Andersen and Horvitz 2007)).

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SET domain methyltransferases contain a ‘switch position’ in their catalytic site that determines the degree of methylation, with bulkier residues able to accommodate mono- and di- but not tri-methylation (Jih et al. 2017). MET-2 and SETDB1 have a bulky

Tryptophan residue in the switch position (Appendix Figure 1B), which suggests that these enzymes favor mono- and di-methylation in the absence of regulatory partners.

Appendix Figure 1. A. Alignment of the amino acid sequences in the SET domain of SETDB1 (red) and G9a (blue) across different species. Worm MET-2 (red) is more similar to SETDB1. ClustalX was used to align sequences. B. The residue highlighted in blue is at the catalytic center of SET domain proteins and determines the degree of methylation. Worm MET-2 and human SETDB1 share a Tryptophan residue (W), whereas other methyltransferases have a Phenyalanine (F).

Analysis of MET-2::GFP hubs in lin-65 and arle-14 mutants

During gastrulation, I observed concentrated hubs of MET-2, LIN-65 and ARLE-14 emerge within nuclei (Chapter 2). The hubs co-localized with puncta of H3K9me2

162

(Chapter 2), suggesting they were sites of active MET-2. Consistent with this idea, MET-

2 hubs were sensitive to loss of lin-65 and to a lesser extent, to arle-14 (Appendix Figure

2). In lin-65 mutants, MET-2 hubs disappeared and the remaining nuclear MET-2 appeared homogenous (Appendix Figure 2B, D). In arle-14 mutants, MET-2 still formed hubs, and these had a similar concentration of MET-2 but were subtly reduced in size

(Appendix Figure 2B-D).

Appendix Figure 2. A. Concentrated protein hubs were defined by using an intensity threshold in ImageJ. The same threshold was applied to wild-type and mutant embryos. B. Single nuclei showing MET-2::GFP hubs in wild-type (green), lin-65 (orange) and arle-14 (yellow) embryos at the gastrula stage (21-50 cell). C. Hub area in wild-type vs. arle-14 mutants (21-50 cell stage). The area of individual MET-2::GFP hubs were measured in ImageJ and averaged for each genotype. Error bars denote standard error of the mean. D. Mean MET-2::GFP intensity in hubs, regions of the nucleus without hubs (non-hub) and the cytosol. Hubs were defined by intensity thresholding. For non-hubs, areas of the nucleus excluding hubs were randomly selected and the intensity measured in ImageJ. For the cytosol, random areas in the cytosol were chosen for intensity measurements. Mean intensity values were averaged for each category in wild-type, lin-65 and arle-14 mutant embryos at the 21-50 cell stage. Error bars denote standard error of the mean.

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Analysis of LIN-65 protein in met-2 and arle-14 mutants

Appendix Figure 3. A. Single sections of whole embryos stained with FLAG and histone antibodies showing distribution of LIN-65::3xFLAG in wild-type vs. met-2, and arle-14 mutants (Scalebar 2 μm). B. Line scans across embryonic cells showing mean LIN-65::3xFLAG intensity in wild- type vs. lin-65 mutants.

LIN-65 is required for nuclear MET-2 accumulation (Chapter 2). I wondered if the same holds true for localization of LIN-65. To analyze LIN-65, I inserted a 3xFLAG tag at the carboxyl terminus of endogenous lin-65 by Crispr (Chapter 2). LIN-65::3xFLAG was nuclear in wild-type embryos and met-2 was required for nuclear localization of LIN-65, whereas arle-14 was dispensable (Appendix Figure 3). These results show that LIN-65 and MET-2 are dependent on each other for nuclear localization.

164

References

Basavapathruni, Aravind, Jodi Gureasko, Margaret Porter Scott, William Hermans, Adarsh Godbole, Peter A. Leland, P. Ann Boriack-Sjodin, Tim J. Wigle, Robert A. Copeland, and Thomas V. Riera. 2016. “Characterization of the Enzymatic Activity of SETDB1 and Its 1:1 Complex with ATF7IP.” Biochemistry 55 (11). American Chemical Society:1645–51. https://doi.org/10.1021/acs.biochem.5b01202.

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