Identification of factors that underly genome-wide transcriptional repression during the development of the Caenorhabditis elegans germline.

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Identification of factors that underly genome-wide transcriptional
repression during the development of the Caenorhabditis
elegans germline.

by
Mezmur D. Belew

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirement for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR BIOLOGY)

August 2023

Copyright 2023 Mezmur D. Belew
ii

Acknowledgments
The completion of the work in this dissertation is not solely to my efforts. I have
several people to thank which I will do in this section. First, I would like to extend my
heartfelt appreciation to both my undergraduate and graduate research advisors, whose
guidance and mentorship pushed me towards pursuing academic research as a career. In Dr.
Engda Hagos’ lab, at Colgate University, I was introduced to the world of biological research
– the highs of successfully developing a new experimental technique and the lows of working
with confusing datasets that seem to contradict themselves. In these early years, I have
learned important lessons such as the patience and time dedication that is needed for
success in research. These are values that I have been and will continue to uphold.

I would also like to thank my doctoral thesis advisor Dr. Matthew Michael for his
mentorship during my time at USC. Matt is a great advisor in many ways. Scientifically, I
admire the way he breaks down big questions into doable experiments in a seemingly simple
way. If there is one trait that I want to learn from him, it would be that. As a mentor, he
demonstrated remarkable patience, which enabled me to get acquainted with a new model
organism and the vast range of new concepts I had to learn to work in his laboratory. He also
pushed me to explore new opportunities that came our way from applying for grants, trying
out internships, and attending conferences. I really have been fortunate to join a group with
as great a PI as Matt.

In addition to my research advisors, I would like to thank the professors at USC that have
been part of my qualifying exam and dissertation committees: Dr. Susan Forsburg, Dr.
iii

Carolyn Phillips, Dr. Peter Calabrese, and Dr. Berenice Benayoun. Their questions and
comments have been quite helpful in designing my proposal and full dissertation
manuscript.

Throughout my time in the Michael Laboratory, I have had the privilege to work with
some great lab-mates. I would like to thank Dr. Matthew Wong who was a senior student
when I joined the laboratory and was responsible for the majority of my technical training.
Through working with him, I was able to get my first publication as a co-author, and he still
gives me advice when I have questions about opportunities after graduate school. Emilie
Chien is another graduate student in our group that has made significant contributions to
the works in this dissertation as we worked together on projects. Her great technical ability
and helpful discussions have helped move the projects forward. I also want to thank Oanh
Huynh for the numerous discussions we've had about experimental designs, which were
always enjoyable and occasionally fruitful. She also came up with our group’s weekly
badminton games which has rapidly grown to include 15 people from the department.

I was also fortunate enough to come in contact with some great people that made the
graduate school experience a good one. I will start with the members of the 2017 cohort:
Joshua Park, Meghan Petrie, Caleb Ghione, Nicole Stuhr, Joe Hale, Yiwei He, and Dan Ma.
These people have been great friends of mine throughout the years. They have been really
helpful at each step of the way from lending ears for complaints and questions about
research, to helping one another during qualifying exam preparations and setting up the
room on the day. Outside of school, our hangouts and birthday dinners have been fun and
iv

memorable. Thank you all for just being great! I would also like to mention new and old
friends like Robel Dagnew, Kaye-Lin Kuphal, Abiy Tekle, and Wilson Ochar and many more
who have celebrated my wins and commiserated with me when I was going through it. All of
these are sincerely appreciated.

Lastly, I want to give the most gratitude to my family. My parents Dawit Belew and Aster
Abate, I value all you have done for me throughout my life. You have worked hard and made
sacrifices to set me up to get to where I am, and I hope I am able to repay a quarter of that in
the times to come. I also appreciate you listening to me every week talking about my
research-related news and getting excited with me. My sister, Mahder Belew, thank you for
being always standing by me and providing your unwavering support whenever I need to
defend my beliefs and actions. Finally, I want to thank my extended family in the US who
have been my immediate support when I needed it. It really took all these people's inputs to
complete this degree and I know I am a lucky person to have them in my life!

v

Table of Contents

Acknowledgements………………………………………………………………………………………………………...ii
List of Figures………………………………………………………………………………………………………………..vi
List of Tables…………………………………………………………………………………………………………………..x
Abstract…………………………………………………………………………………………………………………………xi

Introduction…………………………………………………………………………………………………………………...1
An overview of transcriptional regulation……………………………………………………………..1
The relationship between chromatin organization and transcriptional regulation…..4
The C. elegans germline as a model for transcriptional regulation…………………………..9

Chapter 1: A global chromatin compaction pathway that represses germline gene
expression during starvation in C. elegans……………………………………………………………………...17
Introduction………………………………………………………………………………………………………18
Results and Discussion……………………………………………………………………………………….21
Materials and Methods……………………………………………………………………………………….33
Figures and Figure Legends………………………………………………………………………………..42
Supplementary Materials……………………………………………………………………………………54

Chapter 2: Genome silencing in C. elegans oocytes is triggered by CDK-1 and requires
the topoisomerase II/condensin II axis, the H3K9me pathway, and PIE-1……………………….64
Introduction………………………………………………………………………………………………………65
Results………………………………………………………………………………………………………………69
Discussion………………………………………………………………………………………………………….81
Materials and Methods……………………………………………………………………………………….88
Figures and Figure Legends………………………………………………………………………………..97
Supplementary Figures…………………………………………………………………………………….117

Chapter 3: The TOP-2/condensin II axis silences transcription during germline
specification in C. elegans ………………………… …… ……………………………… …… …………………………...127
Introduction…………………………………………………………………………………………………….128
Results and Discussion……………………………………………………………………………………..131
Materials and Methods……………………………………………………………………………………..140
Figures and Figure Legends………………………………………………………………………………147
Supplementary Figures…………………………………………………………………………………….157

Contributions ……………………………………………………………………………………………………………..163
References………………………………………………………………………………………………………………….164
Introduction references……………………………………………………………………………………164
Chapter 1 references………………………………………………………………………………………...171
Chapter 2 References………………………………………………………………………………………..174
Chapter 3 References………………………………………………………………………………………..180

vi

List of Figures
Introduction
Figure I1: Establishment and development of the C. elegans germline………………………………16

Chapter 1
Figure 1.1: Developmental timing of chromatin compaction in C. elegans primordial
germ cells……………………………………………………………………………………………………………………..42
Figure 1.2: The TOP-2/condensin II axis and heterochromatin pathway are
required for Z2/Z3 chromatin compaction……………………………………………………………………..44
Figure 1.3: A TOP-2/condensin II dependent increase in heterochromatin marks
coincides with chromatin compaction in Z2/Z3………………………………………………………………46
Figure 1.4: Aberrant germline transcription is observed when GCC components
are depleted………………………………………………………………………………………………………………….48
Figure 1.5: GCC pathway and its activator AMPK are required for the nascent
germline to survive nutritional stress……………………………………………………………………………..50
Figure 1.6: Summary of events that comprise a genome activation-repression-
reactivation cycle in the developing C. elegans germline…………………………………………………..52
Figure S1.1: pSer2 antibody reactivity requires the presence of phosphorylated
RNA polymerase II………………………………………………………………………………………………………...54
vii

Figure S1.2: The TOP-2/condensin II axis and heterochromatin pathway are
required for Z2/Z3 chromatin compaction……………………………………………………………………..56
Figure S1.3: Wide-spread H3K9 methylation is observed as chromatin compaction
starts in C. elegans PGCs…………………………………………………………………………………………………58
Figure S1.4: Both H3k9 methyltransferases, MET-2 and SET-25, are needed for the
chromatin compaction in starved L1s……………………………………………………………………………..60

Chapter 2
Figure 2.1: Timing and CDK-9 dependence of the phosphorylation of RNAPIISer2 in
C. elegans proximal oocytes……………………………………………………………………………………………97
Figure 2.2: Aberrant RNAPIIpSer2 signal is observed in proximal oocytes when
TOP-2 and condensin II are depleted………………………………………………………………………………99
Figure 2.3: Aberrant RNAPIIpSer2 signal is observed in proximal oocytes when
CDK-1 is depleted………………………………………………………………………………………………………..101
Figure 2.4: Hyperactivation of CDK by the depletion of WEE-1.3 causes
unscheduled transcriptional silencing………………………………………………………………………….103
Figure 2.5: H3K9me3 signals significantly increase as oocytes become more proximal……105

viii

Figure 2.6: The H3k9 methyltransferases MET-2 and SET-25 are required for
transcriptional repression in proximal oocytes……………………………………………………………..107
Figure 2.7: TOP-2, condensin II and the H3K9 methyltransferases MET-2 and SET-25
are all required for proper bivalent compaction…………………………………………………………….109
Figure 2.8: PIE-1 is required for transcriptional repression in proximal oocytes……………...111
Figure 2.9: PIE-1 is sequestered in the nucleolus prior to silencing…………………………………113
Figure 2.10: Models for how genome silencing occurs in proximal oocytes……………………...115
Figure S2.1: All patterns of RNAPIIpSer2 signal in C. elegans proximal oocytes are
dependent on CDK-9……………………………………………………………………………………………………117
Figure S2.2: TOP-2 and condensin II mediated transcriptional repression in oocytes
is independent of cell-cycle timing………….……………………………………………………………………119
Figure S2.3: CDK-1 levels gradually increase as oocytes become more proximal and
are required for NEB at -1 oocytes………………………………………………………………………………..121
Figure S2.4: Depletion of TOP-2 and condensin II does not alter H3K9me3
deposition in proximal gonads……………………………………………………………………………………..123
Figure S2.5: Representative images for bivalent volume measurements…………………………125

ix

Chapter 3
Figure 3.1: Loss of TOP-2 or CAPG-2 allows for active RNAPII in the P-lineage
of early embryos…………………………………………………………………………………………………………148
Figure 3.2: EGA-GFP gets misexpressed in P4 when TOP-2 and CAPG-2 are depleted………..150
Figure 3.3: EMS specific genes are aberrantly expressed in the P2 cell of 4-cell embryos
after loss of TOP-2 or CAPG-2……………………………………………………………………………………….152
Figure 3.4: The H3K9me heterochromatin pathway is not required for
transcriptional repression in the P lineage……………………………………………………………………154
Figure 3.5: PIE-1 does not require TOP-2 to repress transcription in C. elegans
early embryos……………………………………………………………………………………………………………..156
Figure S3.1: RNAPII-mediated transcription is still repressed in one- and two-cell
embryos after loss of TOP-2 or SET-25………………………………………………………………………….158
Figure S3.2: Control experiments for somatic gene expression in 4-cell embryos…………….160
Figure S3.3: Depletion of MEX-5 and MEX-6 results in PIE-1 presence in all
blastomeres of early embryos………………………………………………………………………………………162

x

List of Tables
Table S1.1: Qualitative measure of signal intensity for immunofluorescence
experiments in this study………………………………………………………………………………………………62
Contributions table……………………………………………………………………………………………………..163

xi

Abstract
The proper transmission of genetic material from generation to generation is an
important aspect of life. In the case of multicellular animals that reproduce sexually,
specialized cell types known as germ cells, are tasked with this. To ensure the fidelity of the
message transmission, germ cells are under constant surveillance for aberrances and employ
multiple mechanisms to avoid them. One mechanism that is well conserved among species
is the tight regulation of transcription in the germ cells during the development of the
organism. To avoid the expression of unwanted genes that may alter the germline fate of
these cells, animals employ gene silencing at specific targets or at a genome-wide level.

In the nematode Caenorhabditis elegans (C. elegans), the development of the germline
is characterized by cycles of genome-wide transcriptional activation and repression. In early
embryos, the P-lineage of cells, that gives rise to the germline, is kept transcriptionally
quiescent by PIE-1. Another round of global transcriptional repression is observed in
primordial germ cells (PGCs) when a newly hatched stage 1 larva (L1) is starved. Unlike the
repression in early embryos, this relies on changes in the chromatin landscape rather than a
single factor facilitating it. Loss of active chromatin marks such as H3K4me, H4K8ac, and
H4K16ac have been reported. Yet, what underlies this change in chromatin architecture and
the extent of the change in chromatin shape still require characterization.

In this study, we delve into the chromatin-based transcriptional repression in
starving L1s. We show that this repression is accompanied by a physical compaction of the
PGC chromatin where the DNA is compacted into bundles that are located at the nuclear
xii

periphery. We also show that this compaction is driven by a novel pathway that entails a
partnership between topoisomerase II (TOP-2), the condensin II complex, and various
heterochromatin-related proteins - such as the H3K9 methyltransferases MET-2 and SET-25
and the H3K9me mark reader proteins CEC-4 and HPL-2. We call this pathway the global
chromatin compaction (GCC) pathway. This pathway is triggered during L1 starvation by the
low-energy sensing proteins AAK-1 and AAK-2 which are homologs of the mammalian
AMPK. Next, we show that the components of the GCC pathway are also required to repress
transcription in maturing proximal oocytes in the adult germline. We also report that the
known transcriptional repressor, PIE-1, also functions in maturing oocytes in addition to P-
lineage cells of early embryos. Finally, we show that the TOP-2/condensin II axis works in
tandem with the PIE-1 to repress transcription in the P-lineage cells of early embryos. All in
all, these studies provide insights into the factors that underly genome wide transcriptional
silencing at different points during the development of C. elegans germline.

1

Introduction
An overview of transcriptional regulation
In 1941, Jacques Monod documented the ability of Escherichia coli (E. coli) to
alternate between sugars as sources of energy based on their availability and gave it the term
“diauxic growth” (Monod, 1949). While their first choice is glucose, if it is not available in the
media, E. coli switch to other sugars like lactose (Lewis, 2011). Work followed to identify the
molecular basis of this ability of bacteria to switch between sugars and it eventually led to
the discovery of operons, a group of genes whose expression is regulated in a concerted
manner and function in a related manner (Jacob and Monod, 1961; Osbourn and Field, 2009).
Jacob and Monod described the detailed workings of the lac operon where the expression of
the lactose metabolizing gene is regulated by a repressor molecule in E. coli (Jacob and
Monod, 1961). This marked the first documentation of molecular control of gene
transcription and gave birth to an enormous field of transcriptional regulation studies within
molecular biology. It is estimated that there are at least 630-700 operons identified in E. coli
(Salgado et al, 2000). Our understanding of their regulation has also become more
sophisticated with the discovery of a diverse host of molecules (proteins, RNAs) regulating
the expression of genes within an operon (Jacob and Monod, 1961; Gollnick, 1994).

Our understanding of the regulation of gene transcription has gone beyond just
operons in the time since Jacob and Monod’s discovery. We now know that genes can be
regulated in multiple ways such as transcription factor binding and chromatin remodeling
such as DNA methylation and various histone modifications (Lorch and Kornberg, 2017;
Weidemüller et al, 2021). Even some RNA species have been shown to play a role in
2

regulating transcription (Guang et al, 2010; Chen and Aravin, 2015). These transcriptional
regulation mechanisms are employed in response to developmental or environmental cues.
For instance, during the development of an organism gene expression is under tight control
to ensure proper cell fate allocation and preservation (Tintori et al, 2016; Moehrle and Paro,
1994). Alternatively, the nutritional status of the organism could influence the expression
levels of genes. Starvation in various organisms ranging from bacteria to mammals results
in altering the levels of expression of several genes involved in metabolism, cell cycle
progression and other cellular processes (Foster, 2005; Klosinska et al, 2011; Wong et al,
2018; Galves et al, 2023).

Based on how many genes are involved, transcriptional regulation schemes can be
classified into two groups: local transcriptional repression and global transcriptional
repression. Local transcriptional regulation operates at the level of an individual or a stretch
of genes either lined up next to each other on a chromosome or occupying the same three-
dimensional region within the nucleus. Transcriptional repression at this level has been
extensively studied and we have several documented examples of it. The lac operon which
has been discussed above is an example of local transcriptional repression. Other examples
of methods of regulating transcription at a local level include histone modifications at
individual genes and small RNA-mediated transcriptional silencing (Cecere, 2021; Parent
and Cahn, 2021). Formation of topologically associated domains (TADs) can also mediate
local transcriptional regulation of genes within chromatin that occupies the same nuclear
region (Gonzalez-Sandoval and Gasser, 2016). TADs, based on their transcriptional activity,
are broadly classified into two groups: type A TADs are domains in which the genes within
3

are prone to transcriptional activity and type B TADs are domains with genes that are
transcriptionally repressed. TADs can also be found interacting with various structures
inside the nucleus such as the nuclear lamina. These special types of TADs are known as
lamina-associated domains (LADs) and are examples of type B TADs (Gonzalez-Sandoval and
Gasser, 2016). This goes in hand with the fact that the nuclear periphery is known to be a
repressive environment and shows how the local environment influences the transcriptional
status of genes (Akhtar and Gasser, 2007).

On the other hand, global transcriptional regulation, as the name suggests, operates
at a genome-wide level. As regulation of gene expression at a genome-wide level is a more
drastic measure than local transcriptional regulation, examples of the former are relatively
rarer. Additionally, the mechanisms that underly global gene regulation are not as clearly
defined. One way to globally regulate transcription is to directly act on the RNA polymerase
enzyme. This is seen in primordial germ cells of Drosophila melanogaster embryos. In these
cells, the protein Pgc is believed to inhibit the phosphorylation of RNA polymerase II
consequently inhibiting transcriptional elongation (Hanyu-Nakamura et al, 2008). In
addition to acting on RNA polymerases, transcription can also be regulated at a global level
through gross changes in chromosomal arrangements. This is seen in organisms of various
degrees of complexity ranging from yeast to humans which showcases how important a role
chromatin plays in regulating the expression of genes (Swygert et al, 2019; Kieffer-Kwon et
al, 2017; Wong et al, 2018). How chromatin rearrangement occurs and its effects on
transcription shall be discussed in more detail in the next section.

4

The relationship between chromatin organization and transcriptional regulation
The term chromatin refers to DNA and its associated proteins known as histone
proteins. In eukaryotes, chromatin can be found in two states: (1) euchromatin – loosely
packed chromatin populated with actively expressed genes and (2) heterochromatin –
densely packed patches of chromatin usually associated with transcriptionally repressed
genes. The definition of these two chromatin states provides an example of the role
chromatin organization plays in regulating gene transcription. In addition to
heterochromatin, which represses transcription constitutively in most cell types, chromatin
reorganization is utilized to control transcriptional activities in a context specific manner
(Swygert et al, 2019; Kieffer-Kwon et al, 2017; Wong et al, 2018; Ishihara et al, 2021). In
addition to transcriptional control, chromatin reorganization plays a role in a multitude of
cellular processes. For instance, during cell division individual chromosomes get condensed
for proper segregation of genetic material into the two daughter cells.

Chromatin reorganization is mediated by distinct sets of enzymes depending on the
cellular context. For instance, heterochromatin formation is marked by known histone
modifications such as the methylations of 9
th
and 27
th
lysine residues of the histone 3 tail
among others. These modifications require methyltransferases such as the SETDB1, EHMT2
(for H3K9 methylation) and the polycomb repressive complex PRC2 (for H3K27
methylation). These histone methylations are bound by multiple reader proteins such as
HP1, which dimerizes to physically compact chromatin (Ahringer and Gasser, 2018). On the
other hand, the chromatin compaction that precedes cell division and the decompaction
afterwards require a separate set of enzymes. Mitotic condensation of chromosomes is
5

mediated by two well characterized factors: the condensin complexes and topoisomerase II.
Work in Xenopus egg extract has shown that after successful replication of DNA,
topoisomerase II is recruited to the chromatin and in turn is required to recruit condensin
complexes. The condensin complexes then facilitate condensation via loop extrusion to form
the mitotic chromosomes that can be faithfully segregated to the daughter cells (Cuvier and
Hirano, 2003; Golfier et al, 2020). Once karyokinesis and cytokinesis are completed,
chromatin compaction needs to be reversed in the daughter cells. This post-mitotic reversal
requires yet another set of enzymes. Mitotic chromosomes are decompacted by a double
hexameric complex formed by the RuvB1 and RuvB2 proteins (Magalska et al, 2014). From
these examples, it is clear that the breadth of different enzymes involved in altering
chromatin states is quite remarkable.

The association between heterochromatin formation and the transcriptional activity
of the genes within it is widely studied (Ahringer and Gasser, 2018; Allshire and Madhani,
2018; McCarthy et al, 2021). Yet, the effect of chromatin remodelers associated with other
cellular processes, such as the mitotic remodelers discussed above, on the transcriptional
status of genes is not as well studied. The following sub-sections will summarize what has
been known so far about the transcriptional regulator functions of the chromatin remodelers
topoisomerase II and the condensin complexes. The remainder of this dissertation will also
provide additional contexts where these proteins regulate gene transcription.

6

The role of topoisomerase II in transcription
The primary function of topoisomerases is to relieve torsional stresses on DNA such
as supercoil formation. Specifically, type II topoisomerases (topoisomerase II) facilitate
double stranded break formation and pass another strand through the breaks to resolve
torsional stresses (Bush et al, 2015). This function of topoisomerase II enables it to play a
role during transcription of genes. Work in budding yeast has shown that the loss of
topoisomerase II results in a global downregulation of genes. And when individual genes are
analyzed in the same study, it was shown that the topoisomerase II is required for the
opening of the promoters by resolving supercoils which enables appropriate transcription
factor binding (Pedersen et al, 2012). Other works have also reported topoisomerase
enrichment at gene promoters and have shown that it is required, along with topoisomerase
I, to efficiently recruit RNA polymerase to the promoters (Sperling et al, 2011).

The double-stranded breaks created by topoisomerase II are typically transient are
resolved by the topoisomerase enzyme itself (Deweese and Osheroff, 2009). In some
instances, the double-stranded breaks made by topoisomerase II may be stable and require
the cell’s DNA damage repair machinery to be resolved (Morimoto et al, 2019; Sasanuma et
al, 2015). These stable breaks have been shown to globally activate transcription in C. elegans
primordial germ cells (PGCs). When a newly hatched C. elegans larva feeds, its PGCs undergo
a large-scale chromatin decompaction. This decompaction requires topoisomerase II-
mediated stable double-stranded breaks, which recruit chromatin decompaction factors like
the RuvBL proteins. This in turn results in the genome wide activation of genes in the PGCs
7

(Wong et al, 2018). These examples highlight the different roles topoisomerase II plays to
regulate transcription.

The role of condensin complexes in transcription
The condensin complexes are multi-subunit complexes first identified as
chromosome organizers in Xenopus egg extracts (Hirano and Mitchison, 1994). Currently,
there are two main condensin complexes well-described in the literature. The first,
condensin I, is located in the cytoplasm and only interacts with chromatin after the nuclear
envelope breaks down during prometaphase. The second condensin complex (condensin II)
is a nuclear complex and has been shown to interact with interphase chromatin (Hirota et al,
2005; Wallace et al, 2015). In terms of subunits, these two condensins contain two common
subunits, SMC2 and SMC4. Condensin I contains 3 additional subunits unique to it: CAP-D2,
CAP-G, CAP-H. Conversely, condensin II contains CAP-D3, CAP-G2 AND CAP-H2 as its unique
subunits (Hirano, 2012).

Condensins have been implicated in chromatin-based regulation of transcription. For
instance, in quiescent budding yeast cells, condensin-mediated large chromosomal
interaction domains (L-CIDs) mediate transcriptional silencing at a global level. These L-CIDs
are formed through loop extrusion, a mechanism of compaction by condensin, of chromatin
up to 12.7 kb in length. Furthermore, the same work shows that this condensin-mediated
chromatin compaction during quiescence is conserved as similar results were observed
when quiescent human foreskin fibroblast (HFF) cells were examined (Swygert et al, 2019).
In addition to specific contexts such as quiescence, condensins have been implicated in the
8

constitutive silencing of pericentromeric repeats and transposable elements. Work done on
Arabidopsis has shown that the condensin subunit SMC4 is required for the appropriate
silencing of pericentromeric transposable elements in concert with cytosine methylases.
This silencing by condensin is not limited to pericentromeric elements in Arabidopsis but
also affects conditionally repressed genes (Wang et al, 2017).

In addition to the two main condensin complexes, some species have additional
condensin complexes that serve specific purposes. Perhaps the most well studied among
these additional condensins is the C. elegans dosage compensation condensin (condensin
I
DC
). Condensin I
DC
differs from condensin I by one subunit as it contains DPY-27 in place of
SMC-4 and is a part of the dosage compensation complex (DCC) (Jans et al, 2009; Ercan,
2015). DCC binds to the X chromosome in hermaphrodite animals (that contain two X
chromosomes) and limits the gene expression by half, resulting in a similar expression of X-
chromosome genes between hermaphrodite animals (XX) and male animals (XO) (Chuang et
al, 1994; Csankovski et al, 2009). DCC binds at sites distributed across the lengths of both X
chromosomes. These sites have been shown to be bound also by the other condensin types,
indicating that condensin I
DC
could play a role in the localization of the DCC on the X
chromosome (Kranz et al, 2013). It has also been reported that DCC does not always bind in
proximity to the genes it represses (Jans et al, 2017). This suggests long range interaction
mediated transcriptional control of genes in the X chromosome. Supporting the long-range
transcriptional regulation, DCC facilitates TAD formation across the X chromosome that
dictates gene expression (Crane et al, 2015). Taken together, the works discussed in this
9

section highlight how condensins reorganize chromatin to modulate gene transcription in
various contexts.

The identification of condensin I
DC
is one example for how useful C. elegans is in the
study of transcriptional regulation. This dissertation also makes use of C. elegans in its
exploration of genome wide transcriptional regulation. The next section will introduce this
powerful model organism and highlight the numerous advantages it provides for
transcriptional regulation studies.

The C. elegans germline as a model for transcriptional regulation
C. elegans are free living nematodes that are commonly found in soil. They are
transparent and can grow to about 1 millimeter in length in their adulthood. They have a
reproductive cycle of 3 days, and their life cycle lasts up to 2 to 3 weeks. C. elegans exist
mostly as hermaphrodites with males accounting for only 0.2 – 0.5% of the population.
Therefore, reproduction takes place mainly through self-fertilization (Hodgkin et al, 1979).
An adult C. elegans lays an average of 300 eggs. The development of the embryos is
characterized by an invariant cell lineage; in other words, each cell division and
differentiation during the development of embryos happens the same way in each and every
individual (Sulston et al, 1983). Once they hatch, C. elegans go through four larval stages,
termed L1 – L4, before reaching adulthood (Meneely et al, 2019).

In early 1970s, molecular biologist Sydney Brenner, who worked primarily with
bacteria and phages, noticed the potential C. elegans had in identifying genes responsible for
10

biological phenomena of interest. He introduced the idea of using these worms for laboratory
studies with a focus on understanding the genetic basis for animal development and
behavior (Meneely et al, 2019). Indeed, the worm possesses multiple qualities that are
appealing to a geneticist like Brenner. First, the fact that majority of reproduction happens
via self-fertilization limits the amount of genetic variation within a population of worms that
stem from a single ancestor. Second, a single worm produces a relatively high brood number
of about 300, which is ideal for statistical analysis. Finally, the short reproductive cycle of
three days makes for quick experimental times.

Since then, the worm has become a mainstay in laboratory studies and has been used
to make seminal discoveries in biology. After their establishment as a laboratory model
organism, the biggest milestone came in 1977: the mapping of C. elegans cell lineage starting
from embryo genesis by Sir John Sulston (Sulston and Horvitz, 1977; Sulston et al, 1983).
Because of these works, we now know what tissue each cell in an early C. elegans embryo
will eventually give rise to. Through this, it was identified that some cells are inevitably
sentenced to death during embryogenesis which led to the discovery of programmed cell
death by Bob Horvitz (Ellis and Horvitz, 1986). The culmination of this was a Nobel prize
being awarded for the two, along with Sydney Brenner for their discoveries regarding
programmed cell death in 2002. The next big step in our understanding of these animals
came when a draft of the C. elegans genome was completed in 1998, making it the first
multicellular organism to have its genome sequenced (C. elegans sequencing consortium,
1998). Through works like these, we currently have a substantial accumulation of knowledge
about C. elegans, which provides an advantage to using them as model organisms.
11

C. elegans are widely used to study gene expression regulation as shown in the
discussion of dosage compensation and condensin I
DC
in the above sections. The discovery
of RNA interference (RNAi) also provided a different, well-conserved mode of gene
regulation. RNAi is a mechanism whereby mRNA of a certain gene is depleted by the
introduction of a double stranded RNA molecule that shares sequence identity with said
mRNA into the organism (Fire et al, 1998). While this mechanism was initially thought of as
a post-transcriptional gene regulation, more recent works have shown that RNAi can
regulate gene expression at the transcriptional level through localized modification of
histones (Mao et al, 2015; Gu et al, 2012; Schwartz-Orbach, 2020; Parent and Cahn, 2021).

Moreover, the C. elegans germline development also provides an ideal platform for
studying transcriptional regulation as the germ cells undergo cycles of genome-wide
transcriptional repression and activation. Work in our laboratory leverages these cycles to
understand the pathways that underly transcriptional regulation at a global level. The
following sub-section provides an overview of the development of the C. elegans germline
(and transcriptional regulation during it) in the hopes of providing an immediate context for
the work I have done during my time in the laboratory.

C. elegans germline development and transcriptional regulation
The assignment of the germline fate in C. elegans occurs as early as the two-cell stage
of embryogenesis where one of the cells becomes destined to give rise to the germline. This
is achieved through an asymmetric segregation of germline fate promoting factors to the
12

posterior side of the zygote, which is thought to be defined by the point of entry of the sperm
(Goldstein and Hird, 1996). The abnormal embryonic partitioning of cytoplasm (par) genes
(par-1, par-2, par-3, and par-6) have been identified as the genes that are required for the
establishment of the asymmetry in a C. elegans zygote. PAR-1 and PAR-2 are localized at the
posterior cortex of the zygote, while PAR-3 and PAR-6 are on the anterior cortex (Kemphues,
2000). This differential distribution of PAR proteins across the cortex promotes the
segregation of germline factors, such as germ granules, to the posterior side of the zygote
(Cheeks et al, 2004; Kemphues, 2000; Figure I1).

Once the first germline precursor is defined, as embryonic development continues, a
series of asymmetric divisions occur to preserve a lineage of single germline precursor cells
known as the P-lineage (Sulston et al, 1983). There are four P-lineage cells, named P1 – P4,
and at the 88-cell stage P4 divides into two primordial germ cells (PGCs) Z2 and Z3. Z2 and
Z3 remain dormant until the embryo hatches and the resulting larva has access to nutrients
(Wang and Seydoux, 2013; Sulston et al, 1983). Throughout the larval development, the PGCs
continue dividing to form the full adult gonad. During the fourth larval stage (L4), the
hermaphrodite C. elegans produces sperm cells and stores it in the spermatheca, then
switches to making oocytes through adulthood (Hirsh et al, 1976). The adult C. elegans
germline is made up of two gonad arms shaped as a ‘U’. The germ cells in each gonad arm are
lined up in order of their development with the most mature oocytes closest to the
spermatheca and immature germ cells further away (Robertson and Lin, 2015; Figure I1).

13

In C. elegans, transcriptional activity in the germline and its precursor cells is tightly
controlled to preserve the germline fate of these cells. In the early embryo, the P-lineage cells
globally repress transcription. PIE-1 (pharynx and intestine in excess) is the factor that
represses transcription. In pie-1 mutant worms, the P-lineage cell in 4-cell embryos (P2)
exhibits aberrant transcription (Seydoux et al, 1996). This leads to the expression of somatic
transcription factors such as SKN-1 and establishes a somatic program in P2. Eventually P2
loses its germline fate, and its descendants adopt the fate of those of its somatic sister cell,
EMS (Mello et al, 1992). These embryos develop into animals that lack a germline, showing
the importance of PIE-1-mediated transcriptional repression during early germline
development (Mello et al, 1992; Nakamura and Seydoux, 2008). However, the mechanism in
which PIE-1 represses transcription is not yet clearly defined. The widely accepted
hypothesis is that PIE-1 prevents the C-terminal domain of RNA polymerase from getting
phosphorylated at the second serine of its heptapeptide repeat. This phosphorylation is
thought to be required for a successful transcriptional elongation (Seydoux et al, 1996;
Palancade and Bensaude, 2003; Nakamura and Seydoux, 2008). But recent work has shown
that the phosphorylation at the second serine is dispensable for embryonic development,
which diminishes its requirement for transcription and suggests that the mechanism of PIE-
1-mediated repression may not be through inhibiting RNA polymerase II serine 2
phosphorylation (Cassart and Yague-Sanz et al, 2023). How PIE-1 represses transcription in
the P-lineage still require more work to be cleared up. Some of the work presented in the
third chapter of this dissertation will provide insights into this.

14

PIE-1 is degraded after the birth of Z2 and Z3 and actively elongating RNA polymerase
II is observed in these cells (Seydoux and Dunn, 1997). At the same time, genome-wide
changes in chromatin marks are observed in the PGCs. H3K4me and H4K8ac, which signify
active chromatin, are lost (Schaner et al, 2003). This suggests a transition to a chromatin-
based transcriptional control in the PGCs. In line with this, work from our group has shown
that PGCs in starved C. elegans L1s are quiescent in terms of transcription and cell cycle.
Their chromatin is also severely condensed into bundles residing at the nuclear periphery.
The induction of PGC transcription upon feeding requires a major decompaction event which
involves the enzyme topoisomerase II (TOP-2) making deliberate double-stranded breaks
on the genomes of the PGCs. These breaks are required to recruit chromatin decompaction
factors such as the RuvBL proteins, RUVB-1/2 (Wong et al, 2018). It is remarkable that C.
elegans go to tremendous lengths of breaking their germline genome to decompact their
chromatin and restart transcription. And it remains a puzzle why they evolved to perform
such a potentially perilous task. A key step in addressing that is understanding how the PGC
chromatin in starved L1s achieves the hyper-compacted structure in the first place. This will
be described in the work presented in the first chapter of this dissertation.

Once an L1 larva hatches and feeds, assuming nutrient is continually provided, its
PGCs remain transcriptionally active as they divide to form the full adult germline. Another
round of global transcription repression is seen in the most proximal maturing oocytes
(Walker et al, 2007). The repression of transcription during oocyte maturation is a
conserved event seen in organisms ranging from simpler metazoans to mammals (Navarro-
Costa et al, 2016; Moore et al, 1974; Schultz et al, 2018). Yet, the factors that underly this
15

silencing have not yet been properly described. One observation that could shed light on how
transcription is repressed is the fact that oocyte chromatin gets significantly compacted as
oocytes approach maturation (Navarro-Costa et al, 2016; Chan et al, 2004). In C. elegans
oocytes, chromatin forms compacted structures known as bivalents. It has been shown that
as oocytes approach maturation condensin II is loaded onto the chromatin and is indeed
involved in the condensation process (Chan et al, 2004). In addition to chromatin
compaction, a recent work has found that PIE-1, the transcriptional repressor that operates
in the P-lineage cells of early embryos, is also present in oocytes (Kim et al, 2021). Whether
chromatin compaction or PIE-1 presence regulates transcription in maturing oocytes has not
yet been shown. Work in the second chapter of this dissertation will elucidate the roles of
chromatin modifying proteins such as condensins and other known transcriptional
repressors proteins play in the silencing of transcription in maturing C. elegans oocytes.

16

Figure I1: Establishment and development of the C. elegans germline.
Green dots signify factors that specify the germline in C. elegans via asymmetric segregation.
See “C. elegans germline development and transcriptional regulation” sub-section of the
introduction for details.

17

Chapter 1: A global chromatin compaction pathway that represses
germline gene expression during starvation in C. elegans.

(Adapted from: Mezmur D. Belew, Emilie Chien, Matthew Wong, W. Matthew
Michael; A global chromatin compaction pathway that represses germline
gene expression during starvation. J Cell Biol 6 September 2021; 220 (9):
e202009197.)

18

Introduction
Sixty years have passed since Jacob and Monod produced a paradigm for the
molecular control of gene expression (Jacob and Monod, 1961), and in that time much has
been learned about how individual genes, or small stretches of genes, are up- and down-
regulated. Far less is known, however, about how gene expression is controlled at larger
scales, for example, at the level of whole genomes. During early development, genome
activation is observed at the maternal-to-zygotic transition (MZT) (Vastenhouw et al., 2019).
In Drosophila, a key factor driving genome activation at the MZT is Zelda, a transcription
factor (TF) which upregulates 100s of genes and appears to function via chromatin
accessibility (Schultz and Harrison, 2019). Genome activation is also observed in human
lymphocytes and is driven by c-Myc, which functions to amplify the low-level of genome
output upon lymphocyte activation (Nie et al., 2012; Percharde et al., 2017).

The studies on Zelda and c-Myc highlight how gene expression is upregulated on a
global basis, and other studies have described how genome output can be globally repressed.
One example of genome repression is found in C. elegans, where the PIE-1 protein interferes
with activation of RNA polymerase II (RNAPII) to repress gene expression during germline
specification in the early embryo (Robert et al., 2015). Loss of PIE-1 allows transcription in
germline progenitors and this, in turn, promotes an exclusively somatic fate for these cells.
Interestingly, while the PIE-1 protein itself is not conserved in other animals, its mechanism
of action is conserved (Robert et al., 2015). A different form of whole-genome repression has
recently been reported for quiescent yeast cells (Swygert et al., 2019). In this system, the
19

chromosome condensation factor condensin remodels chromatin into transcriptionally
repressive loop domains, however it is not yet known if yeast condensin acts alone to
globally repress the genome, or if it is one component of a more complex pathway.

Recent work from our laboratory has examined genome activation, repression, and
re-activation in the C. elegans primordial germ cells Z2 and Z3

(Z2/Z3) (Butuči et al., 2015;
Wong et al., 2018). Z2/Z3 are born during early embryogenesis and, soon after their birth,
the PIE-1 protein is degraded (Mello et al., 1996), allowing transcription to be activated for
the first time in this lineage (Seydoux and Dunn, 1997). Earlier studies had suggested that,
after a period of transcriptional activity in embryonic Z2/Z3, the genome becomes repressed
again (Furuhashi et al, 2010). However, as detailed below, we find that once PIE-1 is
degraded then Z2/Z3 maintain transcriptional competence for the remainder of
embryogenesis and through hatching into L1 larvae. If newly hatched L1 larvae are
maintained in nutrient-free media, however, then transcription is globally repressed,
through a heretofore undefined mechanism (Butuči et al., 2015; Wong et al., 2018). Previous
work from our group has shown that the Z2/Z3 chromatin in starved L1 manifests as tightly
compacted bundles that are positioned exclusively at the nuclear periphery, leaving the
center of the nucleus devoid of chromatin (Wong et al., 2018). Once food is available, a
remarkable series of events unfold that eventually lead to the global reactivation of germline
transcription. In a nutrient-dependent manner, the DNA cleavage enzyme topoisomerase II
(TOP-2) is activated, and it goes on to produce numerous stable DNA double-strand breaks
(DSBs) throughout the genome (Butuči et al., 2015; Wong et al., 2018). The role of the DSBs
20

is to recruit the AAA+ ATPase and chromatin decompacting factor RUVB-1/2 complex to
DNA, and the chromatin is then decompacted. Decompaction leads to a loosening of the
bundles and a relocalization of some chromatin from the periphery to the center. More
importantly, decompaction allows for genome reactivation, and when decompaction fails,
for example after inactivation of TOP-2 or RUVB-1/2, then transcription remains repressed
(Wong et al., 2018). Thus, our work, and work from others, presents a fascinating picture
whereby the nematode germline genome goes through repeated cycles of repression and
activation. The genome is initially repressed via PIE-1 in the early embryo, then activated via
degradation of PIE-1, then repressed again during starvation via the mechanism detailed
below, and finally reactivated when food is present via the DSB-mediated pathway discussed
above.

In the current study, we have focused on how the germline genome is repressed
during starvation. We find that repression occurs through the activity of a global chromatin
compaction (GCC) pathway. By combining genetics with cytology, we identify the
components of the GCC pathway and order their activities within the pathway, thereby
defining a new mechanism of global transcriptional repression.

21

Results and Discussion
Germline chromatin compaction is developmentally regulated and occurs over two stages.
In starved L1s, Z2/Z3 chromatin manifests as globular bundles positioned at the
nuclear periphery (Figure 1.1A, panel “starved L1”; also see Wong et al., 2018). This can be
visualized in living samples using a transgene encoding mCherry-tagged histone H2B (Figure
1.1A), and previous work has shown that when DNA is stained directly in living samples, the
pattern is identical

(Wong et al., 2018). Our first question was: when does germline
chromatin first assume this unusual and highly compacted state? To answer this, we took
single plane images of Z2/Z3 nuclei from living animals of different developmental stages.
We divided the nuclear space into two compartments: an outer compartment (89% of the
surface area) and an inner compartment (11%) (Figure 1.1B) and quantified the fraction of
chromatin that falls into the inner compartment. In germline progenitor cells (P-lineage), the
chromatin was decompacted and diffuse, with ~14% of the chromatin signal located in the
inner compartment (Figures 1.1A&C). At the ~100-cell stage, or ~150 minutes after zygote
cleavage, P4 divides to form Z2/Z3. Shortly after their birth, Z2/Z3 chromatin architecture
was not noticeably different from the P-lineage (Figure 1.1A panel “Z2/Z3 early”; see also
Figure 1.1C), however by the end of gastrulation there was a significant change as the
chromatin became more compacted and the center of the nucleus lost signal (Figures
1.1A&C). This state persisted through the comma and 1.5-fold stages of embryogenesis. We
could not evaluate older embryos due to the onset of muscle twitching that precludes image
capture. However, the pattern was not significantly different in freshly hatched L1s from that
observed in 1.5-fold stage embryos (Figures 1.1A&C). Importantly, however, after overnight
starvation the chromatin became significantly more compacted, with tighter bundles at the
22

periphery and just ~4% of the chromatin signal located in the inner compartment (Figures
1.1A&C). These data reveal a two-stage compaction process for germline chromatin, where
stage I occurs during late embryogenesis and stage II occurs after hatching and during
starvation.

We next wanted to correlate chromatin compaction with the transcriptional status of
Z2/Z3 at various developmental stages. For this, we performed immunofluorescence
microscopy using an antibody that recognizes phospho-Serine 2 (pSer2) within the heptad
repeats of the RNAPII C-terminal domain. The presence of the pSer2 epitope corresponds to
active and elongating RNAPII (Palancade and Bensaude, 2003). Consistent with previous
reports, we found that cells of the P-lineage in early embryos lacked active RNAPII (Seydoux
and Dunn, 1997), whereas the pSer2 epitope was present in Z2/Z3 of all embryonic stages
and was also present in freshly hatched L1s (Figure 1.1D). This suggests that while the P-
lineage is transcriptionally repressed, transcription is activated in Z2/Z3 of early embryos
and remains active through hatching. We note that a previous study had drawn a different
conclusion, in that pSer2 was not detected in Z2/Z3 of late embryos (Furuhashi et al., 2010).
We suspect that this discrepancy lies in the different antibodies used to monitor pSer2. The
previous study used a mouse monoclonal of the IgM subtype (H5), whereas we used a rabbit
monoclonal of the IgG subtype, and thus it may be that the bulkier IgM antibody was unable
to access RNAPII as efficiently as the IgG version used here. We note that the rabbit antibody
used here was validated by us for specificity (Figure S1.1). It thus appears that once PIE-1 is
degraded Z2/Z3 are transcriptionally active through their hatching into L1 larvae. By stark
23

contrast, however, when starved L1s were examined, we observed that pSer2 was no longer
detectable (Figure 1.1D). Thus, during L1 starvation, the chromatin becomes significantly
compacted and transcription is globally repressed.

Both the TOP-2/condensin II axis and the heterochromatin pathway are required for
germline chromatin compaction during starvation.
To learn more about how germline chromatin gets compacted during starvation, we
used RNAi to screen known chromatin compaction factors for a role in this process. Animals
were treated with feeding RNAi against the target gene and their progeny were tested for
Z2/Z3 chromatin compaction after L1 starvation. As shown in Figure 1.2A, and quantified in
Figure 1.2C, depletion of the condensin II subunit CAPG-2 resulted in hypo-compacted
chromatin. The bundles that normally form were diffuse and chromatin was readily
observed in the center of the nucleus, consistent with previous results (Wong et al., 2018).
Similar data were obtained when the KLE-2 subunit of condensin II was targeted (Figures
S1.2A and 1.2C). We next examined TOP-2, as previous work has shown that TOP-2 and
condensin II work collaboratively to resolve sister chromatids and compact chromatin in
preparation for mitosis (Coelho et al., 2003; Cuvier and Hirano, 2003; Shintomi et al., 2015;
Charbin et al., 2014). Like condensin II, depletion of TOP-2 resulted in hypo-compacted
chromatin (Figures S1.2A and 1.2C). We conclude that the TOP-2/condensin II axis is
required for the acute chromatin compaction that occurs during L1 starvation.

24

We next searched for additional components of the compaction mechanism. One
system with a well-known role in compacting chromatin is the heterochromatin pathway
(Ahringer and Gasser, 2018), and recent work in C. elegans has identified CEC-4, an integral
component of the inner nuclear envelope that binds the H3K9 methyl (H3K9me) marks
found in heterochromatin (Gonzalez-Sandoval A et al., 2015). CEC-4 was of interest given
that Z2/Z3 chromatin in starved L1s is packed up against the inner face of the nuclear
envelope. As shown in Figures 1.2B&C, loss of CEC-4 prevented compaction during L1
starvation, as the bundles became diffuse and were no longer organized at the nuclear
periphery. To substantiate this important finding we examined a cec-4 knockout strain,
RB2301, and visualized Z2/Z3 chromatin by direct staining of the DNA with Hoechst 33342
dye in living samples. Relative to the wild-type strain, RB2301 showed disorganized Z2/Z3
chromatin similar to cec-4(RNAi) animals (Figure S1.2B). This effect was germline-specific,
as chromatin architecture in somatic nuclei was not impacted by the depletion of CEC-4
(Figure S1.2C). We also examined another H3K9me “reader” protein, HPL-2 (an HP1
ortholog), and observed that chromatin compaction in starved L1s was perturbed (Figures
S1.2A and 1.2C). Lastly, we wanted to be certain that defects in Z2/Z3 chromatin compaction
are not linked to activation of the RNAi response against an endogenous gene, and this was
confirmed when animals treated with RNAi against mys-1 showed normal compaction
during L1 starvation (Figures S1.2A and 1.2C).

Data shown thus far indicate that Z2/Z3 chromatin is remodeled during both late
embryonic development (stage I compaction) and then again during L1 starvation (stage II
25

compaction), and that components of both the TOP-2/condensin II and heterochromatin
pathways are required for the final compacted state in starved L1s. We next asked if loss of
either TOP-2/condensin II or the heterochromatin pathway would affect stage I compaction,
and we did so by analyzing freshly hatched L1s (termed “new L1s”). As shown in Figures
1.2D-F, loss of both CAPG-2 and TOP-2 had a significant effect on stage I compaction, whereas
depletion of the heterochromatin components CEC-4 or HPL-2 had no effect. It thus appears
that the TOP-2/condensin II axis is active during stage I and the heterochromatin pathway
acts specifically during stage II. This is reasonable, as stage I compaction may simply reflect
the normal chromosome condensation that occurs during G2/prophase as cells prepare for
mitosis, an event known to require TOP-2/condensin II. These data also suggest that stage II
compaction is somehow linked to successful completion of stage I, as otherwise one might
expect a higher degree of stage II compaction in samples depleted for the TOP-2/condensin
II axis. Indeed, as detailed below, this turns out to be the case.

The TOP-2/condensin II axis is required for deposition of H3K9me marks in Z2/Z3
chromatin.
To pursue these observations, we next stained embryos for a canonical
heterochromatin mark, trimethylation of H3 on lysine 9 (H3K9me3) (Ahringer and Gasser,
2018). In early (pre-gastrulation) embryos, H3K9me3 is found in small patches that are
typically positioned at the nuclear periphery (Towbin et al., 2012). This is what we observed,
for both germline progenitor and somatic cells of the early embryo (Figure S1.3A&B). Later
in development, the H3K9me3 signal for early Z2/Z3 (i.e. just after their birth, during early
26

gastrulation) remained similar in intensity to somatic nuclei (Figure S1.3A&B), however by
late gastrulation the H3K9me3 signals in Z2/Z3 became uniformly larger and had spread to
the nuclear interior, whereas the soma remained unchanged (Figure S1.3A&B). The
increased H3K9me3 signal intensity in Z2/Z3 persisted through the remainder of
embryogenesis (Figure S1.3B). Thus, there appears to be a differential deposition of
H3K9me3 in germline versus soma in embryos and this first becomes apparent during late
gastrulation.

We next examined starved L1s for both H3K9me2 and H3K9me3 and observed that
the signals were again much stronger in Z2/Z3 than in nearby somatic nuclei (Figure 1.3A,
panel “N2”). Previous work has shown that H3K9me2/3 deposition in the worm is controlled
by two methyltransferases, MET-2 and SET-25 (Ahringer and Gasser, 2018), and indeed
when mutants for these methyltransferases were examined, we saw a decrease in Z2/Z3
signal intensity (Figure 1.3A). More specifically, we observed that H3K9me2 and me3 were
both compromised in met-2 mutants and me3 was compromised in set-25 mutants. When
both genes were depleted by RNAi then both me2 and me3 marks were attenuated. Both
H3K9me2 and me3 appeared normal in a strain deficient for the SET-32 methyltransferase
(Figure 1.3A). These data show that both MET-2 and SET-25 are required for H3K9me2 and
me3 deposition in Z2/Z3. We next asked if these methyltransferases are required for Z2/Z3
chromatin compaction. RNAi-mediated depletion of either methyltransferase alone, or a
double depletion of both enzymes, resulted in compaction defects in starved L1s, but not
freshly hatched L1s (Figure S1.4). These data show that while the MET-2/SET-25 act during
27

late embryogenesis to hyper-methylate H3K9 in Z2/Z3, the effect of attenuation of these
enzymes is not realized until embryos have hatched and L1s have starved. This is consistent
with our finding that the H3K9me reader proteins CEC-4 and HPL-2 act specifically during
L1 starvation (Figure 1.2).

Our data show that H3K9me marks accumulate in Z2/Z3 during late embryogenesis
and persist within the chromatin through hatching and during L1 starvation, thereby linking
H3K9me deposition to the chromatin compaction that occurs at these times. If deposition of
these marks is truly linked to the compacted state of the chromatin, then, we predicted they
would be removed upon chromatin decompaction during L1 feeding. Indeed, as shown in
Figure 1.3B, upon feeding both H3K9me2 and me3 were rapidly erased, with just a few
patches of peripherally located signals remaining after only one hour. We conclude that, as
Z2/Z3 compact their chromatin in late embryos, H3K9me2/3 marks become intensely
enriched, and during nutrient-dependent decompaction in feeding L1s, the marks are rapidly
lost.

Our data identify two systems, the TOP-2/condensin II axis and the heterochromatin
pathway, as required for germline chromatin compaction. Previous work has linked
condensin and heterochromatin in compaction, in both plants and C. elegans adult germ cells
(Wang et al., 2017; Llères et al., 2017). However, how the two systems are organized to
promote compaction was not known. We, therefore, asked if depletion of capg-2 would
impact the widespread deposition of H3K9me marks that normally occurs on Z2/Z3
28

chromatin in starved L1s. Adult animals were exposed to control or capg-2 RNAi and their
progeny were examined for both H3K9me2 and me3. We observed, surprisingly, that only
very low levels of H3K9me2/3 marks were found in the CAPG-2 depleted samples (Figure
1.3C), and similar results were observed after depletion of TOP-2 (Figure 1.3D). These data
implicate the TOP-2/condensin II axis as acting upstream of MET-2 and SET-25 to promote
H3K9 methylation, but because this was an unanticipated result, we performed a second,
independent, experiment to confirm it. We utilized a transgenic line expressing mKate-
tagged HPL-2 and imaged living samples for HPL-2 signal intensity in Z2/Z3. As shown in
Figures 1.3E&F, for control samples, the HPL-2 signal intensity was similar between Z2/Z3
and somatic nuclei in early embryos but elevated in Z2/Z3 relative to the soma in late
embryos. This is consistent with the onset of heterochromatin assembly occurring in Z2/Z3
of late embryos. By contrast, in capg-2(RNAi) samples, the Z2/Z3 signal intensity, relative to
soma, did not go up nearly as much in late embryos (Figures 1.3E&F), and this is consistent
with a failure to accumulate the H3K9me2/3 marks to which HPL-2 binds. These data place
the TOP-2/condensin II axis upstream of H3K9me deposition during germline chromatin
compaction. This pathway is, to our knowledge, the first to be described that combines the
activities of two well-studied compaction processes into a singular, linear pathway that
compacts chromatin on a global scale. As such we have given this new pathway a name: the
global chromatin compaction (GCC) pathway.

Our data show that GCC is, minimally, a two-stage process that is under both
developmental and metabolic control. Previous work has shown that chromosome
29

condensation in C. elegans embryos is linked to DNA replication, such that condensin II
compacts chromatin in coordination with the replication fork as it is moving along the
chromosome (Sonneville et al., 2015). We propose that stage I compaction in Z2/Z3 amounts
to the replication-dependent and TOP-2/condensin II mediated compaction that occurs in
all cells of the embryo, however, with the additional twist that H3K9me marks are also
deposited, in a germline-specific manner. Interestingly, the H3K9me reader proteins CEC-4
and HPL-2 do not appear to act on the H3K9me marks to promote compaction in late
embryos; rather, they act upon them only when L1s undergo starvation. Thus, the ability of
H3K9me reader proteins to act upon their cognate binding sites on chromatin, and to thereby
promote stage II compaction, is under metabolic control.

GCC is required for transcriptional repression during L1 starvation.
Data in Figure 1.1 show that gene expression in Z2/Z3 is globally repressed during L1
starvation. We, therefore, next asked if transcriptional quiescence is due to GCC. We depleted
GCC components and scored Z2/Z3 in starved L1s for transcription activity by RNAPII pSer2
staining, as in Figure 1.1D. By stark contrast to control samples, depletion of the GCC
pathway components CAPG-2, CEC-4, and MET-2/SET-25 resulted in transcriptionally active
Z2/Z3 nuclei (Figures 1.4A-B). Thus, loss of the GCC pathway leads to RNAPII activation in
starved L1s. We next examined individual genes for misexpression in Z2/Z3, using RNA-FISH
(Choi et al., 2016) to detect the presence of their transcripts in starved L1s. Previous work
from our group has shown that wago-1 and cgh-1 transcripts are absent in starved L1s and
then expressed in Z2/Z3 after L1s have fed

(Wong et al., 2018). We also probed for xnd-1, a
30

germline-specific transcript as a marker for Z2/Z3 (Wong et al., 2108; Mainpal et al., 2015).
As shown in Figure 1.4C, only a small subset of starved wild type L1s showed expression
of wago-1 or cgh-1, consistent with previous results (Wong et al., 2018). However, in
starved cec-4(RNAi) or met-2/set-25(RNAi) L1s, expression of both genes was readily
apparent (Figure 1.4C), with 90-100% of the samples found to contain the transcripts.
Similar results were obtained with two additional genes that we had previously shown to be
upregulated in Z2/Z3 after feeding, ifet-1 and car-1 (Wong et al., 2018). These genes were
mostly silent in control samples but upregulated in top-2(RNAi) L1s (Figure 1.4D). Thus,
across multiple components of the GCC pathway (TOP-2, CAPG-2, CEC-4, and MET-2/SET-
25), we see that depletion leads to both activation of RNAPII and promiscuous gene
expression in the PGCs of starved L1s. We conclude that the GCC pathway represses gene
expression globally in Z2/Z3 during L1 starvation.

The energy sensing kinase AMPK acts as an upstream trigger for GCC.
Having discovered the GCC pathway and shown that it is required for global
transcriptional repression during L1 starvation, we moved on to determine the upstream
trigger for this starvation induced repression. Prior studies have shown that loss of the
energy sensor AMP-activated kinase

(AMPK) results in premature division of Z2 and Z3
during L1 starvation (Demoinet et al., 2017; Fukuyama et al., 2012), a process our group has
shown to require transcriptional activity (Butuči et al., 2015; Wong et al., 2017). Therefore,
given its role in the maintenance of cellular quiescence during starvation, we reasoned that
the upstream trigger for GCC could be AMPK. To test this, we assessed GCC in animals that
31

had been depleted of the AMPK subunits aak-1 and aak-2 using dual RNAi. As shown in
Figure 1.5A, loss of AMPK had a strong impact on stage II compaction, in starved L1s. Stage I
compaction, however, was unaffected (Figure 1.5B). We also asked if depletion of AMPK
would trigger premature activation of RNAPII in starved Z2/Z3 and found that it did (Figure
1.5C). These results place AMPK in the GCC pathway and suggest that AMPK senses the
reduction in energy charge during L1 starvation and, in response, activates GCC to preserve
transcriptional quiescence in the germ cells. We note that one additional phenotype that has
been reported for an aak-1/aak-2 double knockout strain is that of Z2/Z3 cell division during
L1 starvation (Demoinet et al., 2017; Fukuyama et al., 2012). In the aak-1/2 RNAi
experiments shown in Figures 1.2 and 1.5 we looked carefully for >2 PGCs during L1
starvation but we did not encounter this phenotype, and this is likely due to the hypomorphic
effect of RNAi.

In summary, data reported here and elsewhere have revealed a remarkable cycle of
genome-wide activation, repression, and reactivation of transcription in the developing C.
elegans germline (Figure 1.6). The Z2/Z3 genome is activated for the first time after PIE-1 is
degraded, and, in our studies, the genome remains active throughout the remainder of
embryogenesis. It appears, however, that late embryonic Z2/Z3 are indeed preparing for a
global shut down to transcription after hatching. As detailed above, we propose that the TOP-
2/condensin II axis begins to compact the chromatin, likely as a function of DNA replication
(Sonneville et al., 2015), and that TOP-2/condensin II also have an additional function -
promoting the MET-2/SET-25 mediated deposition of H3K9me marks. It is intriguing that
32

H3K9 methylation occurs in earnest at the time of stage I compaction, however these marks
are not needed for stage I compaction and only appear to come into play during stage II
compaction. When L1s starve, we propose that AMPK senses the low energy charge and
responds by triggering stage II compaction, which is mediated by HPL-2 and CEC-4 acting on
the previously deposited H3K9me marks. Stage II compaction is not compatible with
transcription and the genome becomes repressed. Next, when food is available, nutrients
activate TOP-2 to produce stable DSBs, and these are acted upon by RUVB-1/2 to decompact
chromatin and reactivate the genome (Butuči et al., 2015; Wong et al., 2018).

Data presented here define a new mechanism for global regulation of genomic output,
the GCC pathway. While our work has revealed an important new form of transcriptional
regulation, it has also raised fascinating new questions about how whole-genome regulation
is linked to development and metabolism. For example, how does the TOP-2/condensin II
axis promote H3K9me deposition and why is this germline specific? In addition, why do HPL-
2 and CEC-4 only act upon the H3K9me marks during L1 starvation? Surely this is due to
some form of control of HPL-2 and CEC-4 via AMPK, and it will be fascinating to discover the
molecular mechanisms in play.

33

Materials and Methods
Animals
N2 (wild type), WMM1 ([pie-1::gfp::pgl-1 + unc-119(+)]; [(pAA64)pie-
1p::mCherry::his-58 + unc-119(+)] IV), MT13293(met-2(n4256) III), MT17463 (set-
25(n5021) III), VC967 (set-32(ok1457) I), RB2301 (F32E10.2(ok3124) IV), and OH14220
(hpl-2(ot860[hpl-2::mKate2]) III) strains were used in this study. All C. elegans strains were
obtained from CGC or generated in our laboratory. Worms were maintained on 60mm plates
containing nematode growth media (NGM) seeded with the E. coli strain OP50. Worms were
grown at either 20
o
C or 24
o
C and propagated by bleaching or chunking.

Bacterial strains
OP50 bacteria served as the main food source. It was grown in LB media containing
100 ug/ml streptomycin by shaking at 37
o
C overnight. 500 ul of the culture was seeded on
Petri-dishes containing NGM+streptomycin.

Most of our RNAi strains were obtained from the Ahringer library and verified by
Sanger sequencing. An RNAi feeding vector containing the entire hpl-2 (isoform 3) coding
sequence was ordered from Genewiz (South Plainfield, NJ, USA) and was transformed into
HT115 (DE3) cells that were made competent by CaCl2 incubation. Bacteria were then
streaked on LB-agar plates containing 100 ug/ml carbenicillin and 12.5 ug/ml tetracycline
and incubated at 37
o
C overnight. Single colonies were picked and grown in 25ml LB cultures
34

with 100 ug/ml carbenicillin and 12.5 ug/ml tetracycline. 500 ul of this culture was seeded
on 60 mm Petri-dishes containing 5mM IPTG.

Egg preparation
Bleach solution (3.675 ml H2O, 1.2 ml NaOCl, and 0.125 ml NaOH) was prepared.
Adult worms were washed from plates using 5 ml M9 minimal medium (22mM KH2PO4,
22mM Na2HPO4, 85mM NaCl, 2mM MgSO4). Worms were then spun down at 1900 RPM for
1 minute. Excess minimal medium was removed, and the bleach solution was added. Eggs
were extracted by vortexing for 30 seconds and shaking for 1 minute. This was done a total
of 3 times and worms were vortexed one last time. Then the eggs were spun down at 1900
rpm for 1 minute and excess bleach solution was removed and the eggs were washed 3 times
with M9 medium.

RNAi treatment
Feeding RNAi containing NGM plates were prepared as described above. met-2, set-
25, ruvb-1, ruvb-2, and cec-4 feeding vectors were obtained from the Ahringer library while
the hpl-2 feeding vector was produced by us (see above). HT115 cells transformed with an
empty pL4440 vector was used as a negative control for all RNAi experiments. RNAi
conditions used in this study and tests for their efficacy are described below:
top-2 RNAi
35

L1 worms were plated on OP50 plates for the first 53 hours and were then moved to plates
seeded with top-2 RNAi for the remaining 7 hours. Embryonic lethality was observed at
~90%.
met-2/set-25 RNAi
L1s were plated on OP50 plates for 12 hours at 24
o
C. Then worms were transferred to
control and met-2 RNAi plates and were incubated for 24 hrs. Finally, the worms were
transferred from met-2 RNAi plates to set-25 RNAi plates and grown for another 24 hours till
they are gravid adults. When worms are grown at room temperature, the time on OP50 was
extended to 24hrs keeping the RNAi treatment times constant. Once the worms are gravid,
they were bleached and left overnight to hatch. Under these conditions an embryonic
lethality of 15%.
hpl-2 RNAi
L1s were directly plated onto NGM plates seeded with hpl-2 RNAi and were grown for their
whole life until they are gravid adults. Worms were then bleached, and eggs were left to
hatch overnight. Embryonic lethality was ~14 %.
cec-4 RNAi
Worms were grown on cec-4 RNAi plates until gravid after which they were bleached. F1
animals were hatched overnight. Embryonic lethality (~25 %) and defects in Z2/Z3
chromatin compaction was used to ensure RNAi efficacy.

36

capg-2 RNAi
Worms were grown on OP50 plates for the first 44 hours at 24
o
C and were moved to plates
containing capg-2 RNAi for the remaining 16 hours. An embryonic lethality of 80%-90% was
seen with this RNAi treatment.
kle-2 RNAi
Worms were initially grown on food plates and transferred to kle-2 RNAi plates for the final
12 hours till they are gravid adults. An embryonic lethality ranging from 45%-75% was
observed in our experiments.
aak-1/2 RNAi
Worms were grown on plates containing a 1:1 mixture of aak-1 and aak-2 RNAi (referred to
as aak-1/2 RNAi) until they were gravid adults. A 12%-16% embryonic lethality was seen
for these samples.

Antibodies and dilutions
P-granules: Mouse Mab K76 (subtype: IgM) from Developmental Studies Hybridoma
Bank (RRID: AB_531836, DSHB, Iowa City, Iowa) was used at a dilution of 1:20. H3K9me2:
Mouse antibody (subtype: IgG1) from Molecular and Biological Laboratories (MABI0317,
MBL, Woburn, Massachusetts) was used at 1:1000. H3K9me3: Rabbit antibody from Abcam
(ab176916, Abcam, Cambridge, Massachusetts) was used at 1:1000. RNAPII pSer2: Rabbit
antibody from Abcam (ab5095, Abcam, Cambridge, Massachusetts) was used at 1:100.
37

Secondary antibodies: Alexa Fluor conjugated secondary antibodies from Invitrogen
(Thermofisher Scientific, Waltham, Massachusetts) were used at a dilution of 1:200.

Immunofluorescence staining
Animals were first washed with Milli-Q water twice then were spotted on poly-L-
lysine covered slides and let rest for 10 minutes. Coverslips were applied, and slides were
put on dry ice for 25 minutes. Samples were then freeze-cracked by flicking the coverslips
off for permeabilization. Once permeabilized, slides were put in 100% methanol (at -20oC)
for 10 seconds and then fixing solution (0.08M Hepes pH 6.9, 1.6mM MgSO4, 0.8mM EGTA,
3.7% formaldehyde, 1X phosphate-buffered saline) for 10 minutes. After fixing, slides were
washed three times with TBS-T (TBS with 0.1% Tween-20) and were blocked for 2 hours
with TNB blocking solution supplemented with 1:10 normal goat serum (NGS). Primary
antibodies were then applied at the dilutions described above in TNB and slides were
incubated at 4
o
C overnight. The next day, primary antibodies were washed 3 times with TBS
and slides were incubated with secondary antibodies and Hoechst 33342 dye for 2 hours at
room temperature. When two mouse primary antibodies were used in the same experiment,
it was made sure that they were of different subtypes. Subtype-specific secondary antibodies
were then employed so that the distinct antigens could be visualized without signal overlap.
Slides were washed 3 more times with TBS. Mounting medium (50 % glycerol in PBS), and
coverslip were applied and sealed with Cytoseal XYL (Thermofisher).

38

RNA FISH (In Situ Hybridized Chain Reaction)
A kit containing a DNA probe set, DNA hybridized chain reaction (HCR) amplifier
hairpins, and hybridization, wash, and amplification buffers was purchased from Molecular
Instruments (molecularinstruments.com). Genes that were examined were wago-1, cgh-1,
ifet-1, and car-1. A germline-specific gene, xnd-1, was used to mark the primordial germ cells
and Hoechst 33342 dye was used to visualize DNA. Samples were permeabilized using 4%
paraformaldehyde (PFA) and were stored at -80oC for at least one day. Samples were then
treated with Proteinase K (100 ug/ml) for 10 min at 37
o
C and were washed twice with PBS-
T. Samples were treated with 2mg/ml Glycine for 15 minutes on ice and washed twice with
PBS-T. A 1:1 solution of probe hybridization buffer (PHB) and PBS-T was added to the
samples and they were incubated for 5 minutes at room temperature. Samples were then
prehybridized with PHB for 30 minutes at 37
o
C and DNA probes (2 pmol of each) were added
to the samples and were incubated overnight at 37
o
C.

The next day, samples were washed with probe wash buffer (PWB) at 37
o
C and with
5xSSCT at room temperature. Samples were, then, pre-amplified with Amplification buffer
for 30 minutes at room temperature. Probe amplifier hairpins were snap cooled and added
to the sample. Worms were incubated with the hairpins overnight.

On the final day, samples were washed with 5xSSCT at room temperature and
incubated with Hoechst-33342 (1:5000 dilution) for 15 minutes. Finally, samples were
mounted on poly-L-lysine coated slides and imaged.
39

Live-animal imaging
WMM1 and OH14220 samples were treated as needed for the experiments.
To image embryos: Gravid WMM1 adults were bleached and the resulting embryos washed
multiple times with M9 medium. 0.3% agarose pads were made on slides and ~10 ul
embryos were mounted on them. A coverslip was gently applied, and slides were imaged.
To image newly hatched L1s: Gravid WMM1 adults were bleached in the morning. The
resulting embryos were washed three times with M9 medium and were left in M9 for ~5hrs.
Later that afternoon, 0.3% agarose pads were made on slides and ~5 ul of L1s were mounted.
A similar volume of 1M Tetramisole hydrochloride was added to stop the larvae from
moving. Coverslip was gently applied, and slides were imaged.
To image starved L1s: Gravid WMM1 adults were bleached. The resulting embryos were
washed three times with M9 medium and were left overnight in M9 to hatch. The next day,
0.3% agarose pads were made on slides and ~5 ul of L1s were mounted. A similar volume of
1M Tetramisole hydrochloride was added to stop the larvae from moving. Coverslip was
gently applied, and slides were imaged. Identical results were observed when 10 mM sodium
azide was used instead of tetramisole hydrochloride as an anesthetic, see also Wong et al.,
2018.

Hoechst staining of live animals
Gravid adults were bleached, and embryos were left overnight in M9 minimal medium to
hatch. The next day, L1s were spun down at 1900 RPM and transferred to Eppendorf tubes.
40

The remaining M9 was aspirated and worms were incubated with 100 ul of SDS-DTT (804
ul 0.31% SDS; 196 ul of DTT) for 4 minutes. Worms were washed with 1 ml of M9 and then
incubated with 1 ml of 1:5000 Hoechst 33342 dye for 15 minutes. Worms were spun down,
mounted on slides with agar pads and imaged.

Immunofluorescent imaging
All samples were imaged with an Olympus Fluoview FV1000 confocal microscope using
Fluoview Viewer software. Laser intensity was controlled for experiments to achieve
consistency among samples.

Quantification and statistical analysis
GCC assay quantification: Analysis of chromatin compaction was done as described in Wong
et al., 2018. In brief, single-plane images were analyzed using ImageJ. Using the “Oval” tool,
an ellipse was drawn tightly around the chromatin and the total pixel amount was measured
and recorded. Then a smaller ellipse was made by reducing the radii of the original ellipse to
1/3
rd
of its size. This smaller ellipse was positioned at the darkest area at the center of the
nucleus. The number of pixels in that area, which is roughly 11% of that of the original ellipse,
is measured. The fraction of chromatin in the inner 11% was calculated by dividing the pixel
counts in the smaller ellipse by the pixel count of the original ellipse. Data are presented as
the percentage of chromatin that falls into the inner compartment.
41

HPL-2::mKate signal quantification: To quantify HPL-2::mKate signal, whole embryo images
were analyzed using ImageJ. Z2/Z3 were identified in the images by cell size and position.
Once Z2/Z3 are identified, an ellipse was drawn around them using the “Oval” tool. The
number of pixels was measured and recorded. Then, the ellipse was moved to an empty area
to measure the background signal. The actual HPL-2::mKate signal was calculated by
subtracting the background signal from the signal in Z2/Z3. This was also done for a
neighboring somatic cell of comparable size. Then, we calculated the ratio of Z2/Z3 HPL-
2::mKate signal to that of the somatic cell. Data are presented as the fold difference of the
Z2/Z3 signal with respect to the somatic signal.
Statistical analysis: Data were analyzed using either a Student’s t-test or Wilcoxon Rank Sum
test depending on whether the datasets fulfill the requirements for a parametric test or not.
This was determined by using Shapiro-Wilk test for normal distribution and F-test to test for
variance homogeneity of the datasets we were comparing. Differences between two datasets
were considered significant if a p-value of <0.05 was obtained.

Supplementary materials
Figure S1.1 shows the specificity of the antibody used to recognize active RNA polymerase II
in this study. Figure S1.2 contains various data from assays for compaction relevant to Figure
1.2. Figure S1.3 shows that H3K9 hyper-methylation coincides with the onset of chromatin
compaction in Z2/Z3. Figure S1.4 shows more chromatin compaction assays relevant to
Figure 1.2. Table S1.1 contains qualitative signal intensity measures for the
immunofluorescence staining data shown in this study.
42

Figures and Figure Legends

43

Figure 1.1: Developmental timing of chromatin compaction in C. elegans primordial
germ cells.
(A) Germ cell nuclei in living WMM1 embryos of various stages were imaged for
chromatin compaction using an mCherry tagged histone H2B. Scale bar represents a
length of 2 um in this and all other figures. (n = 8)
(B) Schematic showing the two regions of the nucleus used for quantification in
subsequent figures.
(C) Quantification of the data presented in (A). Vertical axis represents the percentage of
pixels in the inner compartment described in (B) and the horizontal axis presents
germ cell nuclei at different stages of development. Error bars represent one standard
deviation. P-values were obtained using the Student’s t-test.
(D) Germ cell nuclei of N2 embryos and larvae at various developmental stages were fixed
and stained for P-granules (white), RNAPII (pSer2) (red) and DNA (blue).
Representative images are shown, with white arrows pointing to P3 in the first panel
and Z2/Z3 in the remaining panels. The number of samples examined ranged from
10 (early embryos) to 22 (starved L1s). See Table S1.1 for a summary of pSer2 signal
intensities across the entire data set.
44

45

Figure 1.2: The TOP-2/condensin II axis and heterochromatin pathway are required
for Z2/Z3 chromatin compaction.
(A) Strain WMM1 was treated with either control or capg-2 RNAi and F1s born from these
animals were starved upon hatching. Z2/Z3 nuclei of the starved L1s were then
imaged for chromatin compaction. (n=20)
(B) Same as (A) except samples were treated with cec-4 RNAi and not capg-2 RNAi.
(C) Chromatin compaction was quantified in starved L1s born from strain WMM1 that
was treated with either capg-2, kle-2, top-2, hpl-2, cec-4 or mys-1 RNAi, as indicated.
Each gene-targeted RNAi was accompanied by a control RNAi, and thus the data are
presented as pairs of control and gene-targeted samples. Error bars represent one
standard deviation. (n=20)
(D) Strain WMM1 was treated with either control or capg-2 RNAi and F1s born from these
animals were examined. Z2/Z3 nuclei of the newly hatched F1 L1s were imaged for
chromatin compaction. (n=20)
(E) Same as (D) except samples were treated with cec-4 RNAi and not capg-2 RNAi.
(F) Chromatin compaction was quantified in newly hatched L1s born from strain WMM1
that was treated with either capg-2, top-2, hpl-2 and cec-4 RNAi. Data are presented
as in (C). Error bars represent one standard deviation. (n=20)

46

47

Figure 1.3: A TOP-2/condensin II dependent increase in heterochromatin marks
coincides with chromatin compaction in Z2/Z3.
(A) Wild-type starved L1s and starved L1s that are defective for methyltransferases
(mutants for met-2, set-25, set-32 and F1s from animals treated with met-2/set-25
double RNAi) were fixed and stained for P-granules (white), H3K9me2 (green),
H3K9me3 (red) and DNA (blue). (n=40) See Table S1.1 for a summary of signal
intensities.
(B) L1s were either starved or fed for a varying amount of time. Samples were then fixed
and stained for P-granules (white), H3K9me3 (red), H3K9me2 (green) and DNA
(blue). Representative images are shown. (n=40) See Table S1.1 for a summary of
signal intensities.
(C) Starved L1s, born from strain N2 treated with either control RNAi or capg-2 RNAi,
were fixed and stained for P-granules (white), H3K9me2 (green), H3K9me3 (red) and
DNA (blue). Representative images are shown. (n=40) See Table S1.1 for a summary
of signal intensities.
(D) Starved L1s, born from strain N2 treated with either control or top-2 RNAi, were fixed
and stained for H3K9me3 (red) and DNA (blue). Representative images are shown.
(n=40) See Table S1.1 for a summary of signal intensities.
(E) Worms that express HPL-2::mKate were optionally treated with control and capg-2
RNAi. Live embryos were extracted and were imaged for HPL-2 signal. The white star
identifies Z2/Z3. Representative images are shown. (n=20)
(F) Quantification of HPL-2::mKate signal from the images taken in (E). Error bars
represent one standard deviation.
48

49

Figure 1.4: Aberrant germline transcription is observed when GCC components are
depleted.
(A) Starved L1s, born from strain N2 that had been treated with either control RNAi
or capg-2 RNAi, were fixed and stained for P-granules (white), DNA (blue), and
RNAPII (pSer2) (green). The number of pSer2 positive samples is shown below.
(n=20)
(B) Starved L1s, born from strain N2 that had been treated with either control RNAi, cec-
4 RNAi or met-2/set25 RNAi, were fixed and stained for P-granules (green), DNA
(blue), and activated RNAPII (pSer2) (red). The number of samples that were positive
for pSer2 signal is shown below. (n=40)
(C) Starved L1s, born from strain N2 that had been treated with either control RNAi, cec-
4 RNAi or met-2/set25 RNAi were used in this experiment. In-situ hybridization chain
reaction (HCR) was performed by probing for wago-1 (red) and cgh-1 (white). DNA
was visualized using Hoechst 33342 dye (blue) and xnd-1 (green) was used to identify
Z2/Z3. Percentage of samples positive for each gene is shown on the right. (n=40)
(D) Starved L1s, born from strain N2 that had been treated with either control or top-
2 RNAi were used. HCR for xnd-1 (green), ifet-1 (red) and car-1 (white) was
performed. DNA was visualized using Hoechst 33342 dye (blue). Percentage of
samples positive for each gene is shown on the right. (n=25)

50

51

Figure 1.5: GCC pathway and its activator AMPK are required for the nascent germline
to survive nutritional stress.
(A) Chromatin compaction was quantified in starved L1s born from strain WMM1 that
was treated with either control or aak-1/ 2 RNAi. Quantification is shown below.
Error bars represent one standard deviation. (n=20)
(B) Chromatin compaction was quantified in newly hatched L1s born from strain WMM1
that was treated with either control or aak-1/ 2 RNAi. Quantification is shown below.
Error bars represent one standard deviation. (n=20)
(C) Starved L1s, born from strain N2 that had been treated with either control RNAi
or aak-1/2 RNAi, were fixed and stained for P-granules (white), DNA (blue), and
RNAPII (pSer2) (red). The number of pSer2 positive samples is shown below. (n=20)

52

53

Figure 1.6: Summary of events that comprise a genome activation-repression-
reactivation cycle in the developing C. elegans germline.
Please see main text for details.

54

Supplementary Materials

55

Figure S1.1: pSer2 antibody reactivity requires the presence of phosphorylated RNA
polymerase II.
Embryos were fixed and stained for P-granules (green), pSer2 (red) and DNA (blue). Both
ama-1 RNAi treatment (which depletes RNAPII) and calf intestine phosphatase treatment
result in the loss of pSer2 signal. Thus, the pSer2 signal is dependent on the presence of
phosphorylated RNAPII.

56

57

Figure S1.2: The TOP-2/condensin II axis and heterochromatin pathway are required
for Z2/Z3 chromatin compaction.
(A) Samples were treated as in Figure 1.2A except the indicated RNAi was used.
(B) Samples were treated as in Figure 1.2D except the indicated RNAi was used.
(C) Starved N2 and RB2301 (cec-4
-/-
) L1s were stained with Hoechst 33342 DNA dye and
live-imaged for Z2/Z3 chromatin compaction. Quantification is shown below. Error
bars represent one standard deviation. (n=20)
(D) Somatic cells in the vicinity of Z2/Z3 in starved L1s, born from strain WMM1 that was
treated with either control or cec-4 RNAi, were imaged for chromatin compaction.
Quantification is shown below. Error bars represent one standard deviation. (n=20)

58

59

Figure S1.3: Wide-spread H3K9 methylation is observed as chromatin compaction
starts in C. elegans PGCs.
(A) C. elegans embryos at different embryonic stages were fixed and stained for P-
granules (green), H3K9me3 (red) and DNA (blue). Representative whole embryo
images are shown.
(B) Same as (A) except only Z2/Z3 (germ) or a neighboring somatic cell (soma) are
shown. The developmental stage of the embryo from which the image was taken is
indicated. (n = 20) See Table S1.1 for a summary of signal intensities.

60

61

Figure S1.4: Both H3k9 methyltransferases, MET-2 and SET-25, are needed for the
chromatin compaction in starved L1s.
Newly hatched and starved L1s born from strain WMM1 treated with either control RNAi,
met-2 RNAi, set-25 RNAi or met-2/set-25 double RNAi were used. Z2/Z3 from the L1s were
imaged for chromatin compaction. Representative images are shown. Compaction was also
quantified and shown below images. Error bars represent one standard deviation. (n=20)

62

Early embryo Late embryo Newly hatched L1s Starved L1s
pSer2 signal detected 0/10 22/25 21/22 4/20
No pSer2 signal detected 10/10 3/25 1/22 16/20
N2 met-2 set-25 set-32 Control RNAi met-2/set-25 RNAi
Strong H3K9me2 signal 40/40 0/40 1/40 39/40 30/40 3/20
Weak H3K9me2 signal 0/40 14/40 37/40 1/40 10/40 28/40
No H3K9me2 signal 0/40 26/40 2/40 0/40 0/40 9/40
N2 met-2 set-25 set-32 Control RNAi met-2/set-25 RNAi
Strong H3K9me3 signal 35/40 0/40 0/20 30/40 34/20 2/40
Weak H3K9me3 signal 5/40 32/40 6/20 10/40 6/40 37/40
No H3K9me3 signal 0/40 8/40 34/20 0/40 0/20 1/40
Starved 1hr feeding 2hr feeding 3hr feeding 4hr feeding
Strong H3K9me2 signal 36/40 5/40 9/20 3/40 3/40
Weak H3K9me2 signal 4/40 35/40 27/40 29/40 23/40
No H3K9me2 signal 0/40 0/40 4/40 8/40 14/40
Starved 1hr feeding 2hr feeding 3hr feeding 4hr feeding
Strong H3K9me3 signal 30/40 7/40 7/40 3/40 5/20
Weak H3K9me3 signal 10/40 33/40 31/40 32/40 24/40
No H3K9me3 signal 0/40 0/40 2/40 5/40 11/40
Control RNAi capg-2 RNAi
Strong H3K9me2 signal 38/40 13/40
Weak H3K9me2 signal 2/40 13/40
No H3K9me2 signal 0/40 14/40
Control RNAi capg-2 RNAi
Strong H3K9me3 signal 34/40 12/40
Weak H3K9me3 signal 6/40 13/40
No H3K9me3 signal 0/40 15/40
Control RNAi top-2 RNAi
Strong H3K9me3 signal 31/40 19/40
Weak H3K9me3 signal 9/40 21/40
No H3K9me3 signal 0/40 0/40
P4 Early Z2/Z3 Gastrula Bean Comma 1.5-fold 2-fold 3-fold
Strong H3K9me3 signal 0/20 4/20 14/20 13/20 17/20 17/20 18/20 17/20
Weak H3K9me3 signal 20/20 16/20 6/20 7/20 3/20 3/20 2/20 3/20
No H3K9me3 signal 0/20 0/20 0/20 0/20 0/20 0/20 0/20 0/20
Figure S1.3B: H3K9me3 staining on N2 embryos at different developmental stages
Figure 1.3D: H3K9me3 staining in starved L1s treated with top-2 RNAi
Figure 1.3C: H3K9me2 and me3 staining in starved L1s treated with capg-2 RNAi
H3K9me2 signal
H3K9me3 signal
H3K9me2 signal
H3K9me3 signal
H3K9me3 signal
Figure 1.3B: H3K9me2 and me3 staining of N2 L1s after 1-4 hrs of feeding
Figure 1.3A: H3k9me2 and me3 staininig in starved L1s defective for heterochromatin formation
Figure 1.1D: pSer2 staining in N2 early/late embryos and starved/fed L1s
H3K9me2 signal
H3K9me3 signal
63

Table S1.1: Qualitative measure of signal intensity for immunofluorescence experiments in
this study

64

Chapter 2: Genome silencing in C. elegans oocytes is triggered by CDK-1
and requires the topoisomerase II/condensin II axis, the H3K9me
pathway, and PIE-1

(Adapted from: Belew MD, Chien E, Wong M, Michael WM. (2023).
Characterization of factors that underlie transcriptional silencing in C. elegans
oocytes. BioRxiv. 2022.08.28.505591)

65

Introduction
Cells can control gene expression at many levels, including individual loci, groups of
genes that perform a common function, or at the level of the entire genome. Whole-genome
control of transcription occurs in many different contexts. For example, cells undergoing
quiescence silence their genomes (Swygert and Tsukiyama, 2019; de Moore and Rando,
2023), as do all proliferating cells as they prepare for mitosis (Palozola et al, 2019; Gonzalez
et al, 2021; Ito and Zaret, 2022). In addition, during germline development, many animals
silence the genome in germline progenitor cells until they have been specified as germline
(Nakamura and Seydoux, 2008; Wang and Seydoux, 2013). Furthermore, a common feature
of gametogenesis is a genome silencing event as gametes complete meiotic prophase
(Schultz et al, 2018; Tora and Vincent, 2021). The genome can also be globally activated, and
examples of this are also found in early development, where many organisms undergo a
zygotic genome activation event as embryos transition from maternal to zygotic control of
embryogenesis (Kobayashi and Tachibana, 2021).

While the concept of whole-genome activation and silencing has been appreciated for
a long time, it is only recently that the molecular mechanisms in play have begun to be
elucidated. One particularly useful model system for the study of whole-genome control of
transcription has been the roundworm C. elegans. Remarkably, during germline
development in the worm, the genome is silenced and then reactivated on at least four
distinct occasions. During early embryogenesis, in one- and two-cell embryos, transcription
is globally repressed via binding of the OMA-1 and OMA-2 proteins to the basal transcription
66

factor TAF-4, as this sequesters TAF-4 in the cytoplasm (Guven-Ozkan et al, 2008). At the
four-cell stage somatic genomes are activated. However, in germline precursor cells, the so-
called P-lineage, transcription remains globally silenced (Mello et al, 1996; Seydoux et al,
1996). Silencing in the P-lineage is controlled by PIE-1, a zinc-finger containing protein
whose mechanism of action during silencing is not yet fully understood. Upon germline
specification, PIE-1 is degraded and germline genome activation occurs in the Z2 and Z3
primordial germ cells (PGCs; Schaner et al, 2003). PGCs remain transcriptionally active
through hatching of the embryo into an L1 larva. Recent work from our group has shown
that if embryos hatch in an environment lacking nutrients, then the PGC genome is silenced
once again (Belew et al, 2021). Starvation-induced genome silencing is triggered by the
energy sensing kinase AMPK and requires the TOP-2/condensin II chromatin compaction
pathway as well as components of the H3K9me/heterochromatin pathway (Belew et al,
2021). In this system, TOP-2/condensin II promotes H3K9me2 and -me3 deposition on
germline chromatin, and thus we named this pathway the Global Chromatin Compaction
(GCC) to reflect the linear organization of the system components. We have also previously
shown that when starved L1s are fed, then the germline genome is reactivated. This requires
the induction of DNA double-strand breaks that serve to promote chromatin decompaction
(Butuči et al, 2015; Wong et al, 2018). Germline chromatin remains transcriptionally active
through the remainder of development and, in hermaphrodites, through oogenesis until the
genome is silenced once again at the end of meiotic prophase (Schisa et al, 2001; Walker et
al, 2007). Thus, in the nematode germline, there are four genome silencing events - in early
embryos by OMA-1/2, in the P-lineage by PIE-1, in starved L1s by the GCC pathway, and in
oocytes via an unknown mechanism.
67

The global repression of transcription during late oogenesis is not unique to C.
elegans. In Drosophila melanogaster, oocytes are developmentally arrested at prophase I and
repress transcription between the fifth and eighth stages of oogenesis, just before their re-
entry into meiosis (Navarro-Costa et al., 2016). Similarly, in mice, primary follicles that
contain prophase I arrested oocytes have been shown to repress transcription just before
the resumption of meiosis (Moore et al, 1974; Schultz et al, 2018; Tora and Vincent, 2021).
This repression persists throughout fertilization until minor ZGA occurs at first cleavage
(Moore and Lintern-Moore, 1974; Abe et al., 2018). The conservation of this theme across
organisms of different complexity suggests that global transcriptional repression in oocytes
is an important feature of the oocyte-to-embryo transition. Despite this conservation,
however, the molecular pathway(s) responsible for repression are still unknown.

In this study, we address the problem of how oocytes in C. elegans shut down
transcription as a function of completing meiotic prophase. The nematode is an ideal system
to address this important question, as the events leading up to oocyte maturation and
fertilization are well described. During germline development in hermaphrodites, animals
first produce sperm and then switch to making oocytes (Schedl, 1997). Oocytes are produced
in an assembly-line like process, where cells exit pachytene and then progress through the
remainder of meiotic prophase within the proximal portion of the tube-shaped gonad
(Greenstein, 2005). Within the proximal gonad, oocytes can be clearly identified by their
position relative to the spermatheca — the oocyte closest is named -1, and the next most
68

proximal oocyte -2, et cetera. Previous work has shown that -4 and -3 oocytes are
transcriptionally active, and then the genome is silenced at the -2 position (Walker et al,
2007; see Figure 2.1A). Other work has shown that, just prior to genome silencing, the
condensin II complex is recruited to chromatin, where it helps to compact chromatin during
the formation of bivalents, a unique chromosome structure that enables the subsequent
meiotic divisions (Chan et al, 2004; see Figure 2.1A). By the -2 position, bivalents have
formed and the genome is silenced. At the -1 position the oocyte receives a signal from sperm
to initiate maturation, and the cells then enter meiotic M-phase (Miller et al., 2001; Figure
2.1A). Here, we show that genome silencing in oocytes is organized by cyclin-dependent
kinase 1 (CDK-1 in C. elegans) and requires the known silencers TOP-2/condensin II, the
H3K9me/heterochromatin pathway, and PIE-1. Loss of any one of these components results
in aberrant RNA polymerase II (RNAPII) activity at the -2 position. Interestingly, we also
report that in oocytes distal to the -2 position, PIE-1 is mainly localized in the nucleolus, and
that at the -2 position the nucleolus dissolves in a TOP-2/condensin II dependent manner.
Our data identify the molecular components for the oocyte genome silencing system and
suggest a model where the nucleolar residency of PIE-1 prevents it from blocking RNAPII
activity until the nucleolus dissolves at the -2 position.

69

Results
RNAPIIpSer2 is dependent on CDK-9 in proximal oocytes.
The goal of this study was to analyze genome silencing during oogenesis in C. elegans.
To monitor transcription, we used an antibody that recognizes the active and elongating
form of RNAPII. This reagent is a rabbit polyclonal antibody termed ab5095, purchased from
Abcam (Waltham, MA), and it detects a phospho-epitope on the second serine within the
carboxy-terminal repeat domain (CTD) of RNAPII (RNAPIIpSer2; Palancade and Bensaude,
2003). In previous work we had validated ab5095 for use in C. elegans via the demonstration
that reactivity depends on the presence of both RNAPII and phosphate, and that ab5095
accurately labels transcriptionally active nuclei in worm embryos and in the PGCs of L1
larvae (Belew et al, 2021). For the current study, we used it to examine the transcriptional
status of oocytes obtained from gonads dissected from adult hermaphrodites. Prior work
that examined proximal oocytes has shown that RNAPIIpSer2 signals decreased starting
with the fourth most proximal oocyte (termed -4) and became undetectable through the two
most proximal oocytes (-2 and -1) (Walker et al., 2007). We, therefore, focused our analysis
on the four most proximal oocytes and found that RNAPIIpSer2 signal was detected
invariably on the chromatin of oocytes at the -4 position (Figure 2.1B, control RNAi panel).
However, as we examined the three most proximal oocytes, we observed several different
patterns for the RNAPIIpSer2 signal: (1) signal exclusively on chromatin, (2) no signal at all,
(3) signal present both on and off chromatin, and (4) a nucleoplasmic RNAPIIpSer2 signal
that mostly excludes chromatin (Figures 2.1B and S2.1A&B). It thus appears that while in
some oocytes RNAPIIpSer2 signal completely disappeared, in others, it was merely removed
70

from the chromatin and was present in the nucleoplasm. This form of nucleoplasmic signal
has also been observed in human cells as they approach mitosis and is explained by RNAPII
coming off the chromatin while maintaining CTD phosphorylations (Hintermair et al, 2016).

In mammals, the kinase that phosphorylates serine 2 within the RNAPII CTD is P-
TEFb, which is composed of the CDK9 kinase and cyclins T1 or T2 (Fujinaga et al, 2023). To
determine if the corresponding kinase in C. elegans, CDK-9, plays a similar role in proximal
oocytes, we used RNAi to deplete the protein and we then stained for RNAPIIpSer2. As shown
in Figures 2.1B and S2.1A&B, exposure to cdk-9 RNAi caused a reduction in all three forms
of RNAPIIpSer2 signal (on chromatin, off chromatin, and both on and off chromatin). This
shows that the off-chromatin signals are due to CDK-9 phosphorylated RNAPII that had been
displaced from chromatin but not yet dephosphorylated. For the remainder of this study, we
only considered nuclei with RNAPIIpSer2 on chromatin as being transcriptionally active, and
the off-chromatin signals were ignored. Interestingly, we also noticed that in more distal
regions of the gonad, for example the pachytene region, RNAPIIpSer2 signal was unchanged
by cdk-9 RNAi (Figure 2.1C). This is consistent with previous work showing that CDK-12 is
the major RNAPIIpSer2 kinase in the mitotic and pachytene regions of the gonad (Bowman
et al, 2013). It thus appears that there is a handoff, from CDK-12 to CDK-9, for generating
RNAPIIpSer2 during oogenesis in C. elegans.

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The TOP-2/condensin II axis controls genome silencing during meiotic prophase.
Our data, together with previous findings (Walker et al, 2007), show that genome
silencing initiates at the -3 position and is largely complete by the -2 position. As detailed
above, there are four genome silencing systems that have been identified thus far in C.
elegans: the TOP-2/condensin II axis (in starved L1 PGCs; Belew et al, 2021), the H3K9me
pathway (in starved L1 PGCs; Belew et al, 2021); the OMA-1/2 proteins (in one- and two-cell
embryos; Guven-Ozkan et al, 2008), and the PIE-1 protein (in the P2, P3, and P4 embryonic
blastomeres; Mello et al, 1996; Seydoux et al, 1996). Which, if any, of these systems might
also be used for genome silencing in oocytes? We know that OMA-1/2 are dispensable for
oocyte silencing, as previous work had directly examined this possibility (Guven-Ozkan et al,
2008). Thus, we turned our attention to the remaining three systems and focused initially on
TOP-2/condensin II. To do so we examined RNAPIIpSer2 in oocytes from animals exposed
to either top-2, capg-2 (which encodes a protein specific for the condensin II complex), or
double top-2/capg-2 RNAi. All three of these treatments impacted the RNAPIIpSer2 pattern
in a similar manner: signal now persisted in -2 oocytes, indicating that genome silencing had
failed (Figure 2.2A&B). To quantify these data, we measured RNAPIIpSer2 signal intensity
present on chromatin within individual oocytes. We normalized these values against signal
intensities present in the pachytene region, as these values do not change upon depletion of
TOP-2 and/or CAPG-2 (data not shown). This was done to account for potential differences
in antibody penetration across the different sample sets (see Methods for a more detailed
explanation). As shown in Figure 2.2B, in the control samples, there was a significant
difference in RNAPIIpSer2 signal intensity between the -3 and -2 positions, reflecting the
genome silencing that occurs in -2 oocytes under normal conditions. Also shown in Figure
72

2.2B is the difference in RNAPIIpSer2 signal intensity between -2 oocytes from control
samples, relative to the samples where TOP-2 and/or CAPG-2 had been depleted. Loss of
TOP-2/CAPG-2 caused a significant increase in RNAPIIpSer2 signal at the -2 position. Based
on these data, we conclude that loss of TOP-2 and/or CAPG-2 impacts genome silencing in -
2 oocytes. Since genome silencing normally occurs in late prophase, one possibility is that
loss of TOP-2/CAPG-2 somehow delays progression through diakinesis, and thus the
persistence of RNAPII transcription in -2 oocytes of top-2/capg-2 depleted samples is an
indirect consequence of defects in prophase timing. This is not the case, however, as when
we stained for the prophase marker phospho-serine 10 of histone H3 (H3pS10; Hendzel et
al, 1997) we saw no difference in the timing of H3pS10 deposition (Figure S2.2). We note
that H3pS10 signal intensity was seemingly increased after top-2/capg-2 RNAi, however in
the absence of an independent source of H3pS10 signal that can be used to normalize signal
intensity across different sample sets it is impossible to know if this is a physiological effect
or an artifact of differential antibody penetration. Nonetheless, it is clear that depletion of
TOP-2/CAPG-2 does not delay prophase timing.

CDK-1 acts upstream of TOP-2/condensin II to trigger genome silencing.
Our data identify a role for TOP-2 and condensin II in the silencing of transcription as
oocytes prepare for maturation and the meiotic divisions. Silencing initiates at the -3
position and is complete by -2, and this is also when condensin II is recruited to oocyte
chromatin (Chan et al, 2004). Previous work has also shown that condensin II activity is
dependent on phosphorylation by CDK1 (Abe et al, 2011), and thus it stands to reason that
73

CDK-1 may also play a role in genome silencing, through regulation of condensin II. To test
this hypothesis, we used RNAi to reduce CDK-1 activity in proximal oocytes and we assessed
the impact on transcriptional activity. As shown in Figures 2.3A&B, cdk-1 RNAi triggered a
significant increase in RNAPIIpSer2 signal intensity in -3, -2, and -1 oocytes, showing that
CDK-1 does indeed control genome silencing. To pursue this further, we next asked the
opposite question – how does elevating CDK-1 activity impact genome silencing? For this we
targeted the CDK-1 inhibitory kinase WEE-1.3 for depletion, as previous work has shown
that WEE-1.3 negatively regulates CDK-1 in nematode oocytes and, further, that CDK-1 is the
sole target of WEE-1.3 (Burrows et al, 2006). Treatment with wee-1.3 RNAi resulted in a
decrease in RNAPIIpSer2 signal intensity across all oocytes spanning the -6 to -2 positions,
showing that genome silencing was happening much earlier than in the control condition
(Figures 2.4A&B). These data posed an important question: is the precocious silencing
observed after wee-1.3 RNAi also dependent on TOP-2/condensin II? To address this, we co-
depleted WEE-1.3 and CAPG-2, and we found that transcriptional activity was now restored
to proximal oocytes (Figures 2.4A&B). Thus, co-depletion with CAPG-2 reverses the effects
of WEE-1.3 depletion alone, and this shows that condensin II is required for the precocious
silencing observed when CDK-1 levels are increased via depletion of WEE-1.3. Based on
these data, we conclude that CDK-1 acts upstream of TOP-2/condensin II, in a positive
manner, to promote genome silencing. Furthermore, our data suggest that the CDK-1 can
activate condensin II for genome compaction and silencing at an activity state below that
needed to trigger nuclear envelope breakdown (NEB) and entry into meiotic M-phase. This,
in turn, suggests that CDK-1 activity rises gradually in the proximal gonad, as opposed to an
abrupt activation in -1 oocytes. This would be consistent with previous work in HeLa cells as
74

well as frog egg extracts showing that CDK-1-cyclin B activity rises gradually during
interphase and prophase (Gavet and Pines, 2010; Maryu and Yang, 2022).To gain additional
evidence for a gradual acquisition of the M-phase state in proximal oocytes we stained them
with MPM-2, an antibody that recognizes mitotic phosphoproteins (Davis et al, 1983). As
shown in Figure S2.3A, MPM-2 antigens are present at low levels in -5 oocytes and they
gradually accumulate as we move proximally in the gonad, such that -1 oocytes show a high
level of MPM-2 reactivity and -3 and -2 oocytes a more intermediate level. These data are
consistent with the idea that CDK-1 activity gradually increases, and that levels of CDK-1 that
are proficient to trigger genome compaction and silencing in -3/-2 oocytes are not yet
sufficient for NEB and entry into meiotic M-phase.

The presence of RNAPIIpSer2 signal at -1 oocytes after cdk-1 RNAi but not after top-
2/capg-2 RNAi also raises the idea that there is an additional transcriptional repression
mechanism in play when CDK-1 activity levels are high enough to trigger meiotic M-phase
entry and NEB. To test this idea, we performed phase-contrast microscopy to assess nuclear
envelope integrity of -1 oocytes in cdk-1 RNAi treated samples. We saw that more cdk-1 RNAi
samples had intact nuclear envelopes than control RNAi treated samples (Figure S2.3B)
establishing a correlation between silencing at -1 oocytes and NEB. When we performed a
similar analysis on wee-1.3 RNAi oocytes (that precociously silence their genome in a top-
2/condensin II dependent manner) we did not see a change in the number of samples with
intact nuclear membrane relative to control RNAi animals (Figure S2.3C). These data show
that the silencing at -1 is tied to NEB progression while the TOP-2/condensin II dependent
75

silencing is not and suggest the existence of an additional silencing mechanism at -1 oocytes
that is distinct from the silencing mechanism discussed so far in this manuscript.

H3K9me3 marks accumulate on oocyte chromatin during genome silencing.
We next examined a role for the H3K9me pathway in oocyte genome silencing. Our
previous work showed that the Z2/Z3 PGCs accumulate H3K9me marks to a significant
extent, relative to neighboring somatic cells, as they prepare for silencing (Belew et al, 2021).
To see if this also occurs on oocyte chromatin we used an antibody, termed ab176916 and
purchased from Abcam (Waltham, MA), that recognizes the tri-methylated form of lysine 9
on histone H3. We have previously validated this antibody with the demonstration that
reactivity is lost in Z2/Z3 of L1 larvae in a strain lacking SET-25, the major H3K9me3
methyltransferase in C. elegans (Belew et al, 2021). We limited the current analysis to
H3K9me3 because available antibodies against H3K9me2 do not recognize their target when
the neighboring serine 10 is phosphorylated (Kimura et al., 2008), and we have seen in
Figure S2.2 that H3pS10 is prominent in proximal oocytes. As shown in Figure 2.5A,
H3K9me3 first appears in -5 oocytes, and then gradually accumulates such that by the -2 and
-1 positions the H3K9me3 and DNA signals overlap extensively. To examine the
reproducibility of this pattern we quantified H3K9me3 signal intensity across 11 gonads and
then compared the values obtained for the -5 to -2 positions to the value obtained for -1
position within a given gonad. As shown in Figure 2.5B, the pattern of gradual accumulation
of H3K9me3 marks as oocytes moved from distal to proximal was indeed highly
reproducible. In C. elegans, H3K9me3 is produced primarily by the SET-25
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methyltransferase (Towbin et al, 2012). Indeed, when H3K9me3 was examined in set-25
mutants and compared to wild type samples, we observed that the H3K9me3 signals were
attenuated (Figure 2.5A). Figure 2.5C shows additional examples of -1 and -2 oocytes for wild
type and set-25 mutants at higher magnification, highlighting the differences in H3K9me3
signal intensity. No such attenuation was observed when met-2 mutants were compared to
wild type (Figure 2.5A), consistent with the MET-2 methyltransferase primarily responsible
for producing H3K9me1 and me2 (Towbin et al, 2012). Our findings are also consistent with
those of Bessler et al, 2010, who observed that H3K9me3 levels in the pachytene region of
the gonad do not change in met-2 mutants, relative to wild type. We conclude that H3K9me3
marks accumulate dramatically on chromatin at the time that oocytes are silencing their
genomes.

The accumulation of H3K9me3 in the oocytes was reminiscent of the H3K9me
spreading we saw during L1 starvation which is dependent on TOP-2 and condensin II
(Belew et al, 2021). Given this, we wished to know if TOP-2 and condensin II also facilitated
the spreading of H3K9me3 in maturing oocytes by staining gonads from top-2/capg-2 RNAi
treated samples for H3K9me3. We saw that there was no difference in H3K9me3 signal
between control and top-2/capg-2 RNAi treated samples (Figure S2.4A and B). Therefore,
unlike in starved L1s, the accumulation of H3K9me3 in maturing oocytes does not require
TOP-2 and condensin II.

77

Both the SET-25 and MET-2 methyltransferases are required for genome silencing in
oocytes.
Having observed SET-25 dependent accumulation of H3K9me3 marks in proximal
oocytes we next asked if SET-25 plays a role in genome silencing. Staining of set-25 mutants
for RNAPIIpSer2 revealed that -2 oocytes contained significantly more RNAPIIpSer2 signal
than did the control samples (Figure 2.6A&B), showing that silencing was attenuated.
Interestingly, we also observed a silencing defect in met-2 mutants (Figure 2.6A&B). Thus,
not only do H3K9me marks accumulate during silencing, but the enzymes responsible for
catalyzing these modifications are also important for genome silencing. We conclude that the
H3K9me pathway globally silences transcription in developing oocytes, as it does in the PGCs
of starved L1s.

Genome silencing is coupled to chromatin compaction in oocytes.
Previous work has shown that condensin II loads on to oocyte chromosomes at the -
3 position and is required for the intense chromatin compaction that occurs as bivalents are
formed (Chan et al, 2004). In starved L1s, genome silencing is coupled to chromatin
compaction (Belew et al, 2021), and thus it was important to monitor compaction in the
oocyte system. For this we turned to a previously utilized strain that carries a transgene
encoding mCherry-tagged histone H2B, which marks chromatin (Wong et al, 2018; Belew et
al, 2021). Living hermaphrodites were immobilized and oocyte chromatin was imaged using
confocal microscopy. We compared control samples to those that been exposed to top-
2/capg-2 RNAi, and we looked at -2 oocytes, which is where genome silencing is occurring.
78

As described in the Methods and Figure S2.5, we measured the volume of the chromatin
masses and found that depletion of TOP-2/CAPG-2 caused a significant increase in volume,
consistent with a defect in compaction, and similar observations were made after set-25 or
met-2 RNAi (Figures 2.7 and S2.5). Thus, as is the case in the PGCs of starved L1s, chromatin
compaction in oocytes is driven by actions of the TOP-2/condensin II axis and components
of the H3K9me pathway. We note that the effects observed here for bivalent compaction
after top-2/capg-2 RNAi are less extreme than those reported by Chan and colleagues, and
this is likely due to differences in how condensin II was inactivated (Chan et al, 2004). In our
experiments, we use a feeding RNAi treatment that targets capg-2. By contrast, Chan and
colleagues combined RNAi with a temperature-sensitive allele, both targeting the hcp-6
subunit of condensin II, thereby using two forms of condensin II inactivation in the same
experiment. As detailed by Chan and colleagues, the effect on bivalent compaction of the
temperature-sensitive hcp-6 (hcp-6
ts
) allele alone, without RNAi, is rather modest.

PIE-1 is required for genome silencing during meiotic prophase in oocytes and localizes to
the nucleolus prior to silencing.
In a final set of experiments, we examined a requirement for PIE-1 in oocyte silencing,
as recent work has shown that PIE-1 is present in the adult gonad (Kim et al, 2021). Figure
2.8A shows that RNAi against pie-1 causes a persistence of transcription in the -2 position,
and quantification shows that RNAPIIpSer2 signal intensity is significantly higher in both -3
and -2 oocytes after pie-1 RNAi, relative to the control samples (Figure 2.8B). Interestingly,
PIE-1 depletion had no effect on bivalent compaction (Figure 2.8C). Thus, like TOP-
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2/condensin II and the H3K9me pathway, PIE-1 is required to repress transcription in -3/-2
oocytes, but unlike these factors its mechanism of action is distinct from chromatin
compaction.

To pursue these findings, we next analyzed PIE-1 localization in oocytes, using a
strain where GFP had been inserted at the endogenous pie-1 locus (Kim et al, 2021). A typical
localization pattern is shown in Figure 2.9A, where we see that PIE-1::GFP is mostly localized
to the nucleus. Furthermore, within the -5 to -3 range of oocytes, it is clear that PIE-1::GFP
accumulated within the nucleolus (Figures 2.9A&B), which can be easily observed using
phase-contrast microscopy (Figure 2.9B). Previous work has shown that as oocytes prepare
for maturation the nucleolus is lost, likely reflecting a shutdown of RNA polymerase I
(RNAPI) transcription (Korčeková et al,2012). This explains why PIE-1::GFP is no longer
predominantly localized to the nucleolus in -2 oocytes, as the nucleolus is undergoing
dissolution (Figures 2.9A&B). Previous work in budding yeast has shown that condensin is
required to remodel rDNA chromatin in preparation for cell division (Freeman et al, 2000).
Given this, we wondered if TOP-2/condensin II is required for nucleolar dissolution in
proximal oocytes. To address this, we used phase-contrast microscopy to image nucleoli, and
we simply measured their area in control and top-2/capg-2 (RNAi) samples. As shown in
Figures 2.9 C&D, for control samples, we observed a significant decrease in nucleolar size in
-2 oocytes, relative to -3 oocytes. This is consistent with the nucleolus undergoing
dissolution in -2 oocytes. When nucleoli were assessed in samples depleted of TOP-2/CAPG-
2, we saw that nucleolar size was significantly increased at both the -3 and -2 positions,
80

relative to controls (Figures 2.9 C&D). Thus, the TOP-2/condensin II axis promotes nucleolar
dissolution in proximal oocytes. We also imaged PIE-1::GFP in samples depleted of TOP-
2/condensin II and observed, as expected, that PIE-1 was localized to the -2 nucleoli that had
resisted dissolution (Figure 2.9E). These data show that the TOP-2/condensin II axis controls
PIE-1::GFP localization, and this likely occurs via TOP-2/condensin II’s ability to promote
nucleolar dissolution. As detailed below in the Discussion, these findings suggest a model for
how TOP-2/condensin II and PIE-1 work together to promote genome silencing in proximal
oocytes.

81

Discussion
The goal of this study was to define the molecular components for genome silencing
in C. elegans oocytes. To monitor transcription, we relied on RNAPIIpSer2 staining, as we and
others have done extensively in the past (see, for example, Seydoux and Dunn, 1997; Walker
et al, 2007; Guven-Ozkan et al, 2008; Cassart and Yague-Sanz et al, 2020; Belew et al, 2021).
One concern with this antibody-based approach is that chromatin compaction in -2 oocytes
may prevent access of the antibody to its target on chromatin, rendering false-negative data.
This is clearly not the case, however, as when we depleted PIE-1 we observed strong
RNAPIIpSer2 signals on chromatin in -2 oocytes, even though compaction occurs normally
under this condition (Figure 2.8). Thus, we consider RNAPIIpSer2 staining to be an accurate
and legitimate method to assess the transcriptional status of oocytes in the worm.

Under normal conditions, we found that RNAPIIpSer2 signal intensity drops
significantly in -2 oocytes relative to the -3 position, and that signals are often undetectable
at -2 (Figure 2.2). This is consistent with previous work (Walker et al, 2007), and thus we
conclude that genome silencing likely initiates at -3 and is largely complete by -2. At the -2
position, all oocytes have intact nuclear envelopes, and thus these cells have yet to enter
meiotic M-phase. This is important as previous work has shown that transcription is
repressed during M-phase (Taylor, 1960; Prescott and Bender, 1962; Parsons and Spencer;
1997), although more recent work has shown that the block is not absolute as a low-level of
transcription can be detected in mitotic cells (Palozola et al, 2017). Nonetheless, it is clear
that M-phase is incompatible with active transcription, and thus it is important to distinguish
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the genome silencing we observe in -2 oocytes from the repression of transcription that
likely occurs during meiotic M-phase in -1 oocytes. Our data support this distinction in
multiple ways. First, depletion of the genome silencing factors TOP-2, CAPG-2, MET-2, SET-
25, and PIE-1 all yield a common phenotype – a persistence of RNAPIIpSer2 signal in -2
oocytes, and loss of the signal in -1 oocytes (Figures 2.2, 2.6, and 2.8). Thus, if loss of active
transcription in -2 oocytes occurs mechanistically the same as in -1 oocytes, then we should
see a persistence of signal in the -1 position also, and we clearly do not. Second, depletion of
CDK-1 results in persistence of RNAPIIpSer2 signals in -2 and -1 oocytes (Figure 2.3). After
exposure to cdk-1 RNAi, -1 oocytes largely retain the nuclear envelope (Figure S2.3B),
consistent with a failure to enter meiotic M-phase. This shows that it is not simply occupancy
of the -1 position that represses transcription; rather, it is M-phase entry that does so. The
finding that transcription is largely repressed during M-phase was made in the early 1960s
however the mechanistic basis is still unknown. A previous study examined hsp70 gene
expression during mitosis and found that three of the four transcription factors (TFs) needed
for hsp70 activation were physically displaced from chromatin at mitosis (Martínez-Balbás
et al, 1995), however how this displacement occurs was not described. It may be that the TFs
are physically displaced by chromosome compaction, or it may be that modification of the
TFs prevents them from binding DNA. Our data shed light on this as we show that M-phase
repression of RNAPIIpSer2 signals is independent of chromosome compaction. We found
that loss of PIE-1 allows compaction, but not RNAPIIpSer2 repression, in -2 oocytes (Figure
2.8). In these same samples, at the -1 position, PIE-1 is no longer needed to suppress
RNAPIIpSer2 as an additional mechanism gets activated. Because there is no further
compaction happening at the -1 position (Chan and Meyer, 2004), this additional mechanism
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is independent of compaction. Indeed, previous work on virus-infected human cells has
shown that M-phase transcriptional repression occurs on nucleosome-free and
uncompacted DNA (Spencer et al, 2000), which is consistent with our data that the M-phase
mechanism is distinct from compaction. In summary, we believe that the PIE-1 data
demonstrate two independent mechanisms for repression at -2 relative to -1. At -2,
chromatin compaction is necessary, but not sufficient, for repression. At -1, repression
occurs on previously compacted chromatin, and thus is mechanistically distinct from what
is happening at -2. Taken together, these data show that genome silencing precedes entry
into meiotic M-phase. Why has the worm evolved a system to block transcription at -2 when
it is going to happen anyway at -1? We propose that active transcription may hamper the
chromosomal remodeling that occurs at -2, and thus that genome silencing at -2 has evolved
to allow proper bivalent formation.

Taking a candidate approach, we found that multiple genome silencing pathways are
operational in -2 oocytes, as loss of the TOP-2/condensin II axis, the H3K9me pathway, and
PIE-1 all impact silencing. We also found that silencing is under cell cycle control, as reducing
CDK-1 activity prevents silencing in -2 oocytes and increasing CDK-1 activity promotes
precocious silencing in oocytes distal to -2 (Figures 2.3 and 2.4). Furthermore, we obtained
evidence that oocytes gradually approach the M-phase state, as MPM-2 antigens accumulate
gradually in the proximal gonad, and not abruptly at the -1 position as one might expect if
CDK-1 is abruptly activated in -1 oocytes (Figure S2.3A). Taken together, these data paint a
picture where CDK-1 activity increases gradually as a function of oocyte position in the
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proximal gonad, and that it is the -2 position where CDK-1 activity crosses a threshold
sufficient to trigger silencing, but not yet sufficient to trigger entry into meiotic M-phase.
Thus, we propose that the timing of genome silencing is controlled by a CDK-1 activity
gradient spanning the proximal gonad (Figure 2.10A).

Our previous work had identified a linear pathway, termed GCC, that silences
germline transcription during L1 starvation (Belew et al, 2021). For GCC, TOP-2/condensin
II acts upstream of the MET-2 and SET-25 methyltransferases to promote H3K9me2 and -
me3 deposition on chromatin (Belew et al, 2021). Interestingly, while TOP-2/condensin II
and SET-25/MET-2 are all required for silencing in oocytes, they are not organized into a
linear pathway as we found that H3K9me3 deposition in oocytes does not require TOP-
2/condensin II (Figure S2.4A and B). H3K9me3 deposition is increased as oocytes approach
the -1 position (Figure 2.5), and this is similar to what happens in late embryos, where
deposition is dramatically increased in Z2/Z3, relative to their somatic counterparts (Belew
et al, 2021). How this increase is regulated, and whether it occurs via spreading of
preexisting marks or via de novo deposition are fascinating questions for future research. As
mentioned above, we were unable to track H3K9me2 marks, but it is interesting to note that
loss of met-2, which is responsible for H3K9me2 deposition, has a stronger effect on silencing
than does loss of set-25, which is responsible for H3K9me3 (Figure 2.6). It thus appears that
both H3K9me2 and -me3 play a role in genome silencing, and another important route for
future research will be to determine which of the H3K9me readers is involved and how they
are acting mechanistically to promote silencing. We note that in mice, it has been appreciated
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for some time that oocytes undergo chromatin compaction concurrent with transcriptional
repression (Zuccotti et al., 1995; Bouniol-Baly et al., 1999; De La Fuente et al., 2004; Schultz
et al, 2018). Interestingly, very recent work has shown that a histone H3.3 chaperone
complex, comprised of the Hira and Cabin1 proteins, promotes H3K9me3 deposition in
chromatin, chromatin compaction, and genome silencing in mouse oocytes (Smith et al.,
2022). These data are consistent with our findings and suggest that a conserved feature of
genome silencing is the H3K9me-mediated compaction of chromatin on a global scale. It will
be of interest to determine if TOP-2 and/or condensin II are also required for genome
silencing in murine oocytes.

Our work also describes a novel function for PIE-1 in oocyte genome silencing. PIE-1
has been well studied in its role of blocking transcription in the P-lineage of early embryos,
however how it does so is still a mystery. Early work suggested a model where PIE-1 binds
to and sequesters cyclin T, a subunit of the CDK9 kinase that phosphorylates RNAPII on
serine 2 within the CTD (Zhang et al, 2003). More recent work, however, has shown that PIE-
1 mutants that fail to bind cyclin T still block transcription in the P-lineage (Ghosh and
Seydoux, 2008) and, furthermore, that the RNAPIIpSer2 mark itself is dispensable for
embryogenesis (Cassart and Yague-Sanz et al., 2020). Thus, it may be that, in the embryo,
PIE-1 acts by interfering with RNAPII serine 5 phosphorylation (Ghosh and Seydoux, 2008),
via an unknown mechanism. If so, this begs the question of why has PIE-1 evolved the ability
to specifically interact with cyclin T? One plausible answer is that PIE-1 targets cyclin T to
block transcription in oocytes. Interestingly, we have shown here that CDK-9, presumably
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acting with cyclin T, is the relevant serine 2 kinase in proximal oocytes, unlike the remainder
of the gonad where CDK-12 is the relevant kinase (Bowman et al, 2013). This finding makes
it intriguing to speculate that the switch from CDK-12 to CDK-9 occurs so that RNAPIIpSer2
can be regulated by PIE-1 in proximal oocytes. Sorting out how PIE-1 is blocking
transcription in oocytes is another important avenue for future research.

Lastly, another important research question raised by our study is how is PIE-1
regulated in the proximal gonad? We have shown that PIE-1-GFP is present in the nuclei of -
5, -4, and -3 oocytes, yet these cells are transcriptionally active (Figure 2.1B and 2.9A).
Importantly, in oocytes distal to -2, PIE-1-GFP is localized predominantly in nucleoli (Figure
2.9). This might explain why these nuclei are competent for transcription despite the
presence of PIE-1, if nucleolar residency prevents PIE-1 from accessing its target(s) for
transcriptional repression. Previous work has shown that the nucleolus dissolves at the -2
position (Korčeková et al, 2012). We have observed this as well and, furthermore, we have
shown that dissolution requires TOP-2/condensin II (Figure 2.9). How might the various
components required for genome silencing in oocytes fit together mechanistically? Our data
support the model shown in Figure 2.10B. We propose that once CDK-1 activity passes a
threshold then TOP-2/condensin II is activated and recruited to chromatin, and this has two
consequences. One, chromatin compaction commences, and this represses transcription,
likely via occlusion of RNAPII and various transcription factors from promoters on the
compacted chromatin. Two, as the rDNA is compacted, RNAPI synthesis is blocked and the
nucleolus dissolves, thereby liberating PIE-1 to block transcription via an unknown
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mechanism. Lastly, independent of TOP-2/condensin II, the methyltransferases targeting
H3K9 are stimulated and H3K9me deposition is hyper-activated, leading to chromatin
compaction and genome silencing. While this model is consistent with our data, there is
clearly much more work needed to establish its accuracy. Future experiments will address
the role of PIE-1 nucleolar residency in its regulation as well as the mechanism by which the
SET-25 and MET-2 methyltransferases are activated and how H3K9me marks accumulate so
dramatically on oocyte chromatin.

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Materials and Methods
C. elegans strains
N2 (wild-type), WMM1 ([pie-1::gfp::pgl-1 + unc-119(+)]; [(pAA64)pie-
1p::mCherry::his-58 + unc-119(+)] IV), MT13293(met-2(n4256) III), MT17463 (set-
25(n5021) III) and WM330 (pie-1(ne4301[pie-1::GFP]) III) strains were used in this study.
Worms were maintained on 60mm plates containing nematode growth media (NGM) seeded
with the E. coli strain OP50 or HT115. Worms were grown at 20
o
C and propagated through
egg preparation (bleaching) every 72 hours.

Bacterial strains
OP50 bacteria served as the primary food source. It was grown in LB media
containing 100 μg/ml streptomycin by shaking at 37
o
C overnight. 500 μl of the culture was
seeded on Petri-dishes containing NGM + streptomycin. HT115 bacteria grown in LB media
containing 100 μg/ml carbenicillin and 12.5 μg/ml tetracycline and seeded on NGM +
carbenicillin + tetracycline plates were also used as a source of food. Our RNAi strains were
obtained from the Ahringer library and verified by Sanger sequencing. Bacteria containing
dsRNA were streaked on LB-agar plates containing 100 μg/ml carbenicillin and 12.5 μg/ml
tetracycline and incubated at 37
o
C overnight. Single colonies were then picked and grown in
25 ml LB cultures with 100 μg/ml carbenicillin and 12.5 μg/ml tetracycline. 500 μl of this
culture was seeded on 60-mm Petri dishes containing 5mM IPTG.

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Egg preparation
Bleach solution containing 3.675 ml H2O, 1.2 NaOCl, and 0.125 ml 10N NaOH was
prepared. Adult worms were washed from plates with 5 ml of M9 minimal medium (22mM
KH2PO4, 22mM Na2HPO4, 85mM NaCl, and 2mM MgSO4). Worms were centrifuged at 1.9
KRPM for 1 minute and the excess medium was removed, then the bleach solution was
added. Eggs were extracted by vortexing for 30 seconds and shaking for 1 minute. This was
done a total of 3 times and worms were vortexed one last time. Then the eggs were spun
down at 1900 rpm for 1 minute and excess bleach solution was removed, and the eggs were
washed 3 times with M9 minimal medium.

RNAi treatment
RNAi containing NGM plates were prepared as described in the “Bacterial strains”
section. For double RNAi treatments, RNAi cultures were mixed at a 1:1 ratio by volume.
HT115 cells transformed with an empty pL4440 vector was used as a negative control. RNAi
conditions used in this study and tests for their efficacy is described below:
cdk-9 RNAi
L1 worms were plated on HT115 food plates for the first 48 hours and were then moved to
plates containing cdk-9 RNAi for the remaining 24 hours. Embryonic lethality in the range of
80% - 85% was observed.
top-2 RNAi
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L1 worms were plated on HT115 food plates for the first 24 hours and were then moved to
plates seeded with top-2 RNAi for the remaining 48 hours. Embryonic lethality was observed
at >90%.
capg-2 RNAi
Worms were grown on HT115 food plates for the first 24 hours and were moved to plates
containing capg-2 RNAi for the remaining 48 hours. An embryonic lethality of 80%-100%
was seen with this RNAi treatment.
top-2/capg-2 double RNAi
Worms were grown on HT115 food plates for the first 24 hours and were transferred to top-
2/capg-2 double RNAi plates for the next 48 hours. Embryonic lethality ranged from 90%-
100% for this RNAi treatment.
cdk-1 RNAi
Worms were grown on HT115 food plates for the first 24 hours and were transferred to cdk-
1 RNAi plates for the next 48 hours. Embryonic lethality ranged from 97%-100% for this
RNAi treatment.
wee-1.3 RNAi
Worms were grown on plates containing wee-1.3 RNAi for the entirety of their life cycle. An
embryonic lethality of approximately 40% was observed. Additionally, a significant
reduction in brood size, and the coalescence of bivalents into one chromatin mass in
proximal oocytes of some samples, were observed for wee-1.3 RNAi worms, as previously
reported (Burrows et al., 2006).
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wee-1.3/capg-2 double RNAi
Worms were grown on plates containing wee-1.3 RNAi for 24 hours then were moved to
plates containing wee-1.3/capg-2 RNAi where they remained for the rest of their life cycle.
An embryonic lethality of approximately80%, and the coalescence of bivalents in proximal
oocytes of some samples were observed.
met-2 RNAi
Worms were grown on plates containing met-2 RNAi for the entirety of their life cycle. Some
of the adult worms were bleached and an L1 chromatin compaction assay was performed on
the resulting larvae to test RNAi efficacy. See Belew et al., 2021, for details on the L1
compaction assay.
set-25 RNAi
Worms were grown on plates containing set-25 RNAi for the entirety of their life cycle. RNAi
efficacy was tested via the same method as for met-2 RNAi.
pie-1 RNAi
Worms were grown on pie-1 RNAi plates for the entirety of their life cycle. An embryonic
lethality of 100% was observed for this RNAi.

Antibodies and dilutions
RNAPIIpSer2: Rabbit antibody from Abcam (ab5095, Waltham, Massachusetts) was
used at a dilution of 1:100. H3pSer10: Rabbit antibody from Rockland Immunochemicals
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(600-401-I74, Pottstown, Pennsylvania) was used at a dilution of 1:500. H3K9me3: Rabbit
antibody from Abcam (ab176916, Waltham, Massachusetts) was used at a dilution of 1:1000.
MPM-2: Mouse antibody (isotype - IgG1) from Sigma-Aldrich (05-368, St. Louis, Missouri)
was used at a dilution of 1:500. Secondary antibodies: Alexa Fluor conjugated secondary
antibodies from Invitrogen (Thermo Fisher Scientific, Waltham, Massachusetts) were used
at a dilution of 1:200.

Immunofluorescence staining
Adult worms were first washed off plates with 10 ml of M9 minimal medium and
rinsed 3 more times. Then, they were centrifuged at 1.9 KRPM and the excess medium was
removed. 20 μl of media containing about 50 worms were spotted on a coverslip and 3 μl of
anesthetic (20mM Sodium Azide and 0.8M Tetramisole hydrochloride) was added to
immobilize them. Worms were dissected using 25Gx5/8 needles (Sigma Aldrich, St. Louis,
Missouri). To release gonads, adult worms were cut twice, once right below their pharyngeal
bulb and once near the tail. The coverslip was then mounted onto poly-L-lysine covered
slides and let rest for 5 minutes. Slides were put on dry ice for 30 minutes. Samples were
then freeze-cracked by flicking the coverslips off for permeabilization.
For RNAPIIpSer2, H3pSer10 and MPM-2 antibody staining experiments, once
samples were permeabilized, slides were put in cold 100% methanol (-20°C) for 2 minutes
and then fixing solution (0.08M HEPES pH 6.9, 1.6mM MgSO4, 0.8mM EGTA, 3.7%
formaldehyde, 1X phosphate-buffered saline) for another 30 minutes. After fixing, slides
were washed three times with TBS-T (TBS with 0.1% Tween-20) and were blocked for 30
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minutes with TNB (containing 100mM Tris-HCl, 200 mM NaCl, and 1% BSA). Primary
antibodies were then applied at the dilutions described above in TNB and slides were
incubated at 4
o
C overnight.
For H3K9me3 staining experiments, permeabilized samples were put in cold 100%
methanol (-20°C) for 10 seconds and then fixing solution (0.08M HEPES pH 6.9, 1.6mM
MgSO4, 0.8mM EGTA, 3.7% formaldehyde, 1X phosphate-buffered saline) for 10 minutes.
After fixing, slides were washed three times with TBS-T (TBS with 0.1% Tween-20) and were
blocked for 2 hours with TNB (containing 100mM Tris-HCl, 200 mM NaCl, and 1% BSA)
supplemented with 10% normal goat serum. Primary antibodies were then applied at the
dilutions described above in TNB supplemented with 10 % goat serum and slides were
incubated at 4
o
C overnight.
On the next day, the slides were washed 3 times with TBS and slides were incubated
with secondary antibodies and Hoechst 33342 dye for 2 hours at room temperature. Slides
were washed 3 times with TBS, mounting medium (50 % glycerol in PBS), and coverslips
were applied and sealed with Cytoseal XYL (Thermo Fisher Scientific, Waltham
Massachusetts).

Live animal imaging
Adult N2, WMM1 and EGW83 worms were collected off plates and were washed 3
times with 10 ml M9 minimal medium. After the last wash, worms were spun down at 1.9
KRPM and excess medium was removed. 0.3% agarose pads were made on slides, and a 10
μl aliquot of adult worms was mounted. 4 μl of anesthetic (20mM Sodium Azide and 0.8M
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Tetramisole hydrochloride) was added to stop the worms from moving. A coverslip was
gently applied, and the slides were imaged.

Immunofluorescent imaging
All slides were imaged using an Olympus Fluoview FV1000 confocal microscope using
Fluoview Viewer software at a magnification of 600x (60x objective and 10x eyepiece
magnifications). Laser intensity was controlled for experiments to achieve consistency
among samples.

Quantification of data
RNAPIIpSer2 signal quantification
For each oocyte nucleus, two images were taken: one of the Hoechst-stained DNA and
one of the RNAPIIpSer2 signal. Images were analyzed using ImageJ. An outline was drawn
using the polygon selection tool around the Hoechst-stained area to mark the space occupied
by DNA. The region of interest was copied and pasted to the RNAPIIpSer2 signal image and
the raw integrated density (the sum of the values of the pixels in the selection) was
measured. The raw integrated density was then normalized by the area to get a measure we
called “signal density”. To account for possible variability in signal intensity due to different
degrees of antibody penetration from sample to sample, each oocyte's signal density was
normalized to an average signal density from 5 pachytene nuclei found in the same image.
The final normalized signal densities for the oocytes are presented. For each condition, data
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were collected from two independently performed experimental replicates and the data
were then pooled for statistical analysis and presentation.
H3K9me3 signal quantification
Signal densities for each oocyte were calculated similarly to what is described above
in the RNAPIIpSer2 signal quantification. For data on Figure 2.5, H3K9me3 signal of each
oocyte in the -2 to -5 positions was normalized to the most proximal (-1) oocyte and the
ratios are presented as percentages. For data presented in Figure S2.4 signal densities of
control and top-2/capg-2 RNAi samples were shown side by side for comparison.
H3pSer10 signal quantification
Signal densities for each oocyte were calculated similarly to what is described above
in the RNAPIIpSer2 signal quantification. Signal densities were not normalized to pachytene
nuclei since pachytene nuclei either did not harbor any signal. (for H3pSer10 staining) or the
signals were affected by our RNAi treatments (for H3K9me3 staining) which precluded us
from using them for unbiased normalization.
Bivalent volume quantification
Z-stacks of oocyte nuclei were acquired. For each condition, 5 nuclei were analyzed
and every bivalent within those nuclei that were visually distinguishable from one another
was included in our analysis. On average 4 bivalents per nucleus were analyzed using ImageJ.
The scale was set to 69nm per pixel. To measure the volume of a bivalent, a polygon was
tightly drawn around it on each stack the bivalent appears in. The areas of these polygons
were measured and summed up. Finally, the sum was multiplied by the distance between
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each stack to calculate an approximation of the bivalent’s volume. Averages of the volumes
of all bivalents analyzed were presented.
Nucleolar size measurement
Images were captured of focal planes corresponding to the maximum diameter of the
nucleolus in living oocytes for each oocyte position. Diameters were measured using ImageJ
and then converted to area.

Statistical analysis
Prior to performing any statistical test, data was tested whether it was parametric or
not. To do so, the Shapiro-Wilk test was used to test for normal distribution and F-test was
used to test for variance homogeneity of the datasets we were comparing. Data were then
analyzed using a student’s t-test or Wilcoxon Rank Sum test depending on whether the
datasets fulfill the requirements for a parametric test or not. Differences between any two
datasets were considered statistically significant if a P-value of <0.05 was obtained.

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Figures and Figure Legends

98

Figure 2.1: Timing and CDK-9 dependence of the phosphorylation of RNAPIISer2 in C.
elegans proximal oocytes.
A. Schematic summarizing transcriptional activity and chromatin compaction in the
four most proximal oocytes. See introduction for details.
B. N2s were treated with either control or cdk-9 RNAi. Dissected gonads from these
animals were fixed and stained for DNA (blue) and RNAPIIpSer2 (red). Depletion of
CDK-9 results in the loss of RNAPIIpSer2 signal in proximal oocytes. Scale bar
represents a length of 2 µm.
C. Pachytene nuclei from the same animals in (B) were fixed and stained for DNA (blue)
and RNAPIIpSer2 (red). Unlike proximal oocytes cdk-9 RNAi does not alter
RNAPIIpSer2 signal in pachytene nuclei. Scale bar represents a length of 2 µm.

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100

Figure 2.2: Aberrant RNAPIIpSer2 signal is observed in proximal oocytes when TOP-
2 and condensin II are depleted.
A. N2 animals were treated with control, top-2, capg-2, or top-2/capg-2 double RNAi.
Gonads were dissected from these animals and were fixed and stained for DNA (blue)
and RNAPIIpSer2 (red). Each panel series (-3, -2, -1, and pachytene) represent nuclei
from the same gonad. Individual depletion of TOP-2, CAPG-2, and their co-depletion
results in persistent RNAPIIpSer2 signal on the chromatin of -3 and -2 oocytes. Scale
bar represents a length of 2 µm.
B. Quantification of data presented in (A). RNAPIIpSer2 signal was measured using
ImageJ. The average ratio of normalized raw integrated density is plotted on the y-
axis. Between 10 and 22 gonads were analyzed per condition and data were collected
across multiple independently performed replicates. For control RNAi 22 gonads
across 5 replicates were analyzed, for top-2 RNAi 15 gonads were analyzed across 2
replicates, for capg-2 RNAi it was 20 gonads across 2 replicates and for top-2/capg-2
RNAi it was 10 gonads across a single replicate. Significance was measured using
student’s t-test or Wilcoxon Rank Sum test. **p<0.01; *p<0.05. In control animals, the
ratio of signal density decreases significantly between -3 and -2 oocyte positions. Loss
of TOP-2 or CAPG-2 results in persistent RNAPIIpSer2 signal in -2 oocytes. Co-
depletion of TOP-2 and CAPG-2 produces a stronger phenotype, with persistent
RNAPIIpSer2 signal in -3 and -2 oocytes.

101

102

Figure 2.3: Aberrant RNAPIIpSer2 signal is observed in proximal oocytes when CDK-
1 is depleted.
A. N2 animals were treated with control or cdk-1 RNAi. Gonads were dissected from
these animals and were fixed and stained for DNA (blue) and RNAPIIpSer2 (red). Loss
of CDK-1 results in RNAPIIpSer2 signal persisting into the -2 and -1 oocyte positions.
B. Quantification of data presented in (A). RNAPIIpSer2 signal was measured using
ImageJ. The average ratio of normalized raw integrated density is plotted on the y-
axis. 19 samples were analyzed for each RNAi treatment over 2 independent
replicates. Significance was measured using student’s t-test or Wilcoxon Rank Sum
test. **p<0.01; *p<0.05. Depletion of CDK-1 resulted in a significant increase in on-
chromatin RNAPIIpSer2 signal in the three most proximal oocytes.

103

104

Figure 2.4: Hyperactivation of CDK by the depletion of WEE-1.3 causes unscheduled
transcriptional silencing.
A. N2 animals were treated with either control, wee-1.3, or wee-1.3/capg-2 double RNAi.
Gonads were dissected from these animals and were fixed and stained for DNA (blue)
and RNAPIIpSer2 (red). Depletion of WEE-1.3 results in the loss of on-chromatin
RNAPIIpSer2 signals in the more distal (-6 to -4 position) oocytes. Co-depletion using
wee-1.3/capg-2 RNAi reverses the loss of RNAPIIpSer2 signal. Scale bars represent a
length of 2 µm.
B. Quantification of data presented in (A). RNAPIIpSer2 signal was measured using
ImageJ. The average ratio of normalized raw integrated density is plotted on the y-
axis. 20 samples were analyzed for each RNAi treatment over 2 independently
performed replicates. Significance was measured using student’s t-test or Wilcoxon
Rank Sum test. **p<0.01; *p<0.05. Depletion of WEE-1.3 resulted in a decrease in
RNAPIIpSer2 signal from -6 to -2 oocyte positions. Co-depletion of WEE-1.3 and
CAPG-2 resulted in restored RNAPIIpSer2 signal in -6 to -3 oocyte positions.

105

106

Figure 2.5: H3K9me3 signals significantly increase as oocytes become more
proximal.
A. Wild-type, met-2 mutant, and set-25 mutant gonads were dissected, fixed, and stained
for DNA (blue) and H3K9me3 (red). In N2s, an increase in H3K9me3 signal is
observed at the -3 to -1 positions in comparison to more distal oocytes and pachytene
nuclei from the same animal. Loss of MET-2 does not affect the H3K9me3
accumulation pattern. Depletion of SET-25 results in loss of H3K9me3 signal in
proximal oocytes. Scale bar represents a length of 2 µm.
B. Quantification of data for wild type samples in (A). H3K9me3 signals for each oocyte
in -5 to -2 positions relative to the most proximal oocyte are presented. 20 samples
were analyzed over 2 independent replicates. H3K9me3 signal accumulates on
chromatin in the more proximal oocyte positions.
C. Additional examples of -2 and -1 oocytes stained as in part (A). Either wild type (N2)
or set-25 mutant oocytes are shown. Note that the set-25 samples have only trace
amounts of H3K9me3 on the chromatin, relative to the wild-type sample.

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108

Figure 2.6: The H3K9 methyltransferases MET-2 and SET-25 are required for
transcriptional repression in proximal oocytes.
A. Gonads from wild-type, met-2 mutant, and set-25 mutant animals were dissected,
fixed, and stained for DNA (blue) and RNAPIIpSer2 (red). Mutations of either met-2
or set-25 results in persistent RNAPIIpSer2 signal on chromatin, compared to wild-
type animals. Scale bar represents a length of 2 µm.
B. Quantification of data presented in (A). RNAPIIpSer2 signal was measured using
ImageJ. The average ratio of normalized raw integrated density is plotted on the y-
axis. 10 to 20 samples were analyzed for each condition over 2 independent
replicates. 15 wild-type samples were analyzed while 10 met-2 mutant and 20 set-25
mutants were analyzed. Significance was measured using student’s t-test or
Wilcoxon Rank Sum test. **p<0.01; *p<0.05. Loss of MET-2 resulted in a significant
persistence of RNAPIIpSer2 signal on chromatin in -2 and -1 oocytes. Loss of SET-25
led to a significant persistence of RNAPIIpSer2 signal on chromatin in -2 oocytes.

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110

Figure 2.7: TOP-2, condensin II and the H3K9 methyltransferases MET-2 and SET-25
are all required for proper bivalent compaction.
Living proximal oocytes, treated with control, top-2/capg-2, met-2, or set-25 RNAi, were
imaged for chromatin compaction using a strain harboring mCherry-tagged histone H2B.
Bivalent volume in the -2 oocyte was measured using ImageJ. The average bivalent volume
is plotted on the y-axis. 5 oocyte nuclei (with an average of 4 bivalents per nucleus) were
analyzed for each RNAi treatment. Significance was measured using student’s t-test or
Wilcoxon Rank Sum test. **p<0.01; *p<0.05. Exposure to top-2/capg-2, met-2 or set-25 RNAi
treatments results in significantly larger bivalents.

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112

Figure 2.8: PIE-1 is required for transcriptional repression in proximal oocytes.
A. N2 animals were treated with either control or pie-1 RNAi. Gonads were dissected
from these animals and were fixed and stained for DNA (blue) and RNAPIIpSer2 (red).
Depletion of PIE-1 results in persisting RNAPIIpSer2 signal in proximal oocytes.
B. Quantification of data presented in (A). RNAPIIpSer2 signal was measured using
ImageJ. The average ratio of normalized raw integrated density is plotted on the y-
axis. 20 samples were analyzed for each RNAi treatment over 2 independent
replicates. Significance was measured using student’s t-test or Wilcoxon Rank Sum
test. **p<0.01; *p<0.05. Oocytes from the pie-1 RNAi treatment had significantly
increased RNAPIIpSer2 signal at the -3 and -2 positions compared to control RNAi.
C. Living proximal oocytes treated with control or pie-1 RNAi were imaged for
chromatin compaction using a strain harboring mCherry-tagged histone H2B.
Bivalent volume in the -2 oocyte was measured using ImageJ. The average bivalent
volume is plotted on the y-axis. 5 oocyte nuclei (with an average of 4 bivalents per
nucleus) were analyzed for each RNAi treatment. Significance was measured using
student’s t-test or Wilcoxon Rank Sum test. Exposure to pie-1 RNAi treatment did not
affect bivalent volume.

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Figure 2.9: PIE-1 is sequestered in the nucleolus prior to silencing.
A. The normal localization pattern of PIE-1::GFP in oocytes of live WM330 worms. From
the -5 to -3 oocyte positions, PIE-1::GFP is sequestered within the nucleolus. Oocyte
position is numbered. Scale bar represents a length of 10 µm.
B. A wild-type localization pattern of PIE-1::GFP at the -3 oocyte position. PIE-1::GFP
appears within the nucleolus. Nucleolus is indicated by the white circle. Scale bar
represents a length of 10 µm.
C. WM330 animals were treated with control or top-2/capg-2 double RNAi. Live adults
were imaged at the -3 and -2 oocyte positions. Scale bar represents a length of 5 µm.
D. Quantification of data presented in (C). Nucleolar size was measured using ImageJ
and is plotted on the y-axis. Significance was measured using student’s t-test.
**p<0.01; *p<0.05. Exposure to top-2/capg-2 treatment results in significantly larger
nucleoli.
E. Live adults were imaged for PIE-1::GFP after treatment with control or top-2/capg-2
double RNAi. Loss of top-2/capg-2 results in PIE-1::GFP remaining sequestered in the
nucleolus at the -2 oocyte position. Scale bar represents a length of 5 µm.

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116

Figure 2.10: Models for how genome silencing occurs in proximal oocytes.
A. A CDK-1 activity gradient allows for genome silencing at the -2 position to occur prior
to entry into meiotic M-phase at the -1 position.
B. A proposed pathway whereby CDK-1 promotes TOP-2/condensin II mediated
compaction of rDNA, and this in turn promotes nucleolar dissolution and
relocalization of PIE-1 to the nucleoplasm where it can block transcription.

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Supplementary Figures

118

Figure S2.1: All patterns of RNAPIIpSer2 signal in C. elegans proximal oocytes are
dependent on CDK-9.
A. The different patterns of RNAPIIpSer2 (red) signal on and/or off DNA (blue) observed
in N2s treated either control or cdk-9 RNAi. Depletion of CDK-9 results in the loss of
RNAPIIpSer2 signal in proximal oocytes regardless of RNAPIIpSer2 signal
localization. Scale bar represents a length of 2 µm.
B. Visualization of data presented in (A). N2s treated with cdk-9 RNAi showed reduced
RNAPIIpSer2 signal in all forms. The number of samples analyzed over 2 independent
replicates is presented below the charts for each oocyte position.

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120

Figure S2.2: TOP-2 and condensin II mediated transcriptional repression in oocytes
is independent of cell-cycle timing.
A. N2 animals were treated with control or top-2/capg-2 double RNAi. Gonads from
young adults were dissected, fixed, and stained for DNA (blue) and H3pSer10 (green).
Exposure to top-2/capg-2 RNAi does not affect the timing of H3pSer10. Scale bar
represents a length of 2 µm.
B. Quantification of data presented in (A). 11 control RNAi and 10 top-2/capg-2 RNAi
samples were analyzed for each RNAi treatment. There was no significant difference
in H3pSer10 signal after TOP-2 and CAPG-2 co-depletion. Signal density was not
normalized.

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122

Figure S2.3: CDK-1 levels gradually increase as oocytes become more proximal and
are required for NEB at -1 oocytes.
A. N2 gonads were dissected, fixed, and stained for DNA (blue) and MPM-2 (white). An
increase in MPM-2 signal is observed at the proximal oocyte positions in comparison
to more distal oocytes. Scale bar represents a length of 2 µm.
B. N2 samples were treated with either control or cdk-1 RNAi. Nuclear membrane of -1
oocytes were evaluated using live phase-contrast microscopy. 20 samples were
analyzed over two replicates and quantifications are presented below each
representative image. cdk-1 RNAi resulted in a higher number of samples with intact
nuclear envelope when compared to control RNAi treatment.
C. N2 samples were treated with either control or wee-1.3 RNAi. Nuclear envelope
integrity was evaluated like in (B) for oocytes in -1 to -6 positions. 5 gonads were
analyzed for each treatment and the quantifications are presented below each
representative image. Treatment with wee-1.3 RNAi did not alter the progression of
NEB in proximal oocytes.

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Figure S2.4: Depletion of TOP-2 and condensin II does not alter H3K9me3 deposition
in proximal gonads.
A. Gonads from N2 adults treated with either control or top-2/capg-2 RNAi were
dissected, fixed, and stained for DNA (blue) and H3K9me3 (red). Treatment with top-
2/capg-2 RNAi did not alter the deposition of H3K9me3 in proximal gonads. Scale bar
represents a length of 2 µm.
B. Quantification of signal density for the data presented in (A). 10 samples were
analyzed for each treatment. H3K9me3 signal remained the same when top-2/capg-
2 RNAi treated samples were compared with those treated with control RNAi.

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126

Figure S2.5: Representative images for bivalent volume measurements.
Z-stacks were taken from living oocytes at the -2 position in animals treated with control,
top-2/capg-2, met-2, set-25, and pie-1 RNAi. Shown here are representative images of a
bivalent volume measurement from each RNAi treatment. See methods for details on
bivalent volume measurement. Scale bar represents a length of 2 µm.

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Chapter 3: The TOP-2/condensin II axis silences transcription during
germline specification in C. elegans

(Adapted from: Mezmur D. Belew, Matthew Wong, Emilie Chien, W. Matthew
Michael. (2022). The TOP-2-condensin II axis silences transcription during
germline specification in C. elegans. bioRxiv 2022.08.30.505898)

128

Introduction
Just like individual genes, the transcriptional output of entire genomes can be
regulated in a signal-mediated manner. In fact, genome repression and activation are
recurring events during germline development and gametogenesis in the roundworm C.
elegans. In meiotic prophase, for example, oocytes globally repress transcription as they
prepare for the meiotic divisions (Walker et al, 2007, Belew and Chien et al, 2023). Known
transcriptional repressors, TOP-2, condensin II, heterochromatin pathway and PIE-1 are all
involved in the silencing of oocytes during meiotic maturation (Belew et al, 2021; Seydoux
et al, 1996; Belew and Chien et al, 2023). After fertilization, transcription remains repressed,
but at this time repression is mediated by the OMA-1/2 proteins that bind to and inactivate
the essential transcription factor TAF-4 (Guven-Ozkan et al, 2010). The OMA system blocks
transcription through the first two mitotic divisions and then the proteins are rapidly
degraded. Loss of OMA-1 and 2 activates transcription in three of the four cells in the 4-cell
embryo, with the holdout being the germline precursor P2. P2 remains transcriptionally
repressed via the action of PIE-1, and this is the case for the remainder of P-lineage
development (Mello et al, 1996; Seydoux et al, 1996; Seydoux and Dunn, 1997).

Upon division of the P4 cell, to form the Z2/Z3 primordial germ cells (PGCs), PIE-1 is
rapidly degraded, and the germline robustly activates transcription for the first time (Mello
et al, 1996; Schaner et al, 2003). Z2/Z3 are transcriptionally active for the remainder of
embryogenesis, however if the embryo hatches into a nutrient-free environment then the
PGC genome is rapidly silenced, and this requires the GCC pathway (Belew et al, 2021). Upon
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feeding, Z2/Z3 activate transcription again, in a manner that depends on the deposition of
DNA breaks in the genome that allow the chromatin to decompact (Butuči et al, 2015; Wong
et al, 2018). Thus, in summary, cells within the germline system shut down transcription
during meiotic prophase by a host of repressors (TOP-2, condensin II, heterochromatin
pathway and PIE-1). Then, upon fertilization, repression is maintained first by OMA and then
PIE-1. Transcription is activated in Z2/Z3 and then shut down again in starved L1s via GCC,
and nutrients then allow for reactivation. What is remarkable about this system is the
diversity of mechanisms used to repress transcription, from meiotic prophase repressors to
OMA to PIE-1 and then the GCC pathway.

In this study we focused on transcriptional repression in early embryonic P-cells. The
mechanism by which PIE-1 inhibits transcription is still poorly understood but has been
linked to the phosphorylation of RNA polymerase II (RNAPII) on serines 2 and 5 within its
carboxyl-terminal domain (CTD; Robert et al, 2015; Wang and Seydoux, 2013). The RNAPII
CTD contains numerous copies of the heptapeptide repeat sequence Y1S2P3T4S5P6S7, with
humans containing 52 copies and nematodes 42 copies (Phatnani and Greenleaf, 2006).
During the transcription cycle, RNAPII arrives at the promoter in an unphosphorylated state
and is then phosphorylated on serine 5 to allow promoter clearance. Phosphorylation on
serine 2 follows, and this has been linked to an active, elongating state of RNAPII (
Zaborowska et al, 2016). PIE-1 blocks both serine 5 and serine 2 phosphorylation, and
genetic experiments using separation-of-function mutants have shown that the ability to
block serine 5 phosphorylation is critical to suppressing transcription in the P lineage,
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whereas the ability to block serine 2 phosphorylation is dispensable (Ghosh and Seydoux,
2008). More recent work has shown that the phosphorylation of serine 2 on RNAPII CTD is
dispensable for embryonic development (Cassart and Yague-Sanz et al, 2020). This softens
the link between active transcription and RNAPII serine 2 phosphorylation and supports the
idea that PIE-1 represses transcription by acting on Serine 5 phosphorylation. How PIE-1
blocks serine 5 phosphorylation, mechanistically, and if it works alone to repress
transcription in the P-lineage is not understood. In this work we asked if the GCC pathway
also plays a role in silencing transcription in P-cells, as it does in the PGCs of starved L1s
(Belew et al, 2021). We find that the GCC per se is not required, as we found no role for the
H3K9me pathway in silencing P-cell transcription. Interestingly, however, we do reveal a
role for the TOP-2/condensin II axis in P-cell silencing. We report that loss of TOP-
2/condensin II allows for actively elongating RNAPII in P-cells. We also show that somatic
genes get misexpressed in the P-lineage when TOP-2 and condensin II are lost. Finally, we
present data suggesting that PIE-1 and TOP-2 act separately from one another to repress
transcription in the P-lineage. All in all, these data show that PIE-1 is not the whole story for
silencing in the P-cell lineage and set the stage for detailed mechanistic investigations into
how TOP-2/condensin II and PIE-1 block transcription in the P-lineage.

131

Results and Discussion
Loss of TOP-2 and condensin II function results in active RNAPII in the P lineage of
early embryos.
Previous work from our laboratory had defined the GCC pathway as a signal-
mediated genome silencing system (Belew et al, 2021), and thus we were interested in
determining if GCC was active in the P-cells of early embryos, which are known to be
transcriptionally repressed (Robert et al, 2015, Wang and Seydoux, 2013). To do so, we
stained early embryos with an antibody that detects a phospho-serine 2 on the RNAPII CTD
(referred to as RNAPIIpSer2 for the remainder of this study). The presence of RNAPIIpSer2
signal has been used as a marker for actively elongating RNA polymerase II (Palancade and
Bensaude, 2003), and we and others have used this approach extensively to identify
transcriptionally active nuclei within a variety of C. elegans tissues (Seydoux and Dunn, 1997;
Walker et al, 2007; Wong et al, 2018; Belew et al, 2021; Belew and Chien et al, 2023). The
antibody that we are using, a rabbit polyclonal that recognizes RNAPIIpSer2, has previously
been validated by us for use in C. elegans embryos (Belew et al, 2021). We first examined
zygotes and 2-cell embryos that had been exposed to either control RNAi or RNAi against
two different components of the GCC pathway, top-2 or set-25. As expected, the control RNAi
samples did not show any RNAPIIpSer2 signal, in either zygotes or 2-cell samples, and the
same was true of the top-2 RNAi and set-25 RNAi samples (Figure S3.1). We conclude that
the GCC pathway is not involved in transcriptional repression in one- and two-cell embryos.

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It is at the 4-cell stage that the somatic precursor cells ABa, ABp, and EMS activate zygotic
transcription (Seydoux and Fire, 1994). The remaining cell, P2, remains transcriptionally
repressed due to the presence of PIE-1. We stained 4-cell embryos for RNAPIIpSer2 and for
the control samples the expected pattern emerged – signal could be seen in ABa, ABp, and
EMS, but not P2 (Figure 3.1). Interestingly, in samples exposed to either top-2 or capg-2 RNAi,
we could now detect RNAPIIpSer2 in P2, and the same was true of samples exposed to pie-1
RNAi (Figure 3.1). We next examined the remainder of the P-lineage (P3 and P4) and in both
cases we could detect active transcription after depletion of either TOP-2, CAPG-2, or PIE-1
(Figure 3.1). Based on these data, we conclude that TOP-2 and condensin II are dispensable
for transcriptional repression in P0 and P1, but for P2, P3, and P4 they are required to silence
gene expression.

A genome activation reporter gets misexpressed in the P-lineage when TOP-2 and
condensin II are depleted.
Once we found evidence for active transcription in the P-lineage when TOP-2 and
condensin II are depleted, we then sought to strengthen our findings by identifying specific
gene misexpressions. Prior works have used a C. elegans strain containing GFP reporter
fused to vet-4 promoter to mark early embryonic genome activation (Tocchini et al, 2014;
Fassnacht et al, 2018). For the remainder of this manuscript, this reporter is referred to as
EGA-GFP. It has been shown that vet-4 expression is restricted to somatic cells of early
embryos with P2 lacking any vet-4 transcript in 4-cell embryos (Seydoux et al, 1996). Using
the EGA-GFP strain, we asked if the depletion of TOP-2 and condensin II resulted in the
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misexpression of the GFP reporter in P-lineage cells. In our hands, we saw no EGA-GFP signal
until P4 was born although vet-4 mRNAs have been seen in as early as 4/8-cell embryos
(Seydoux et al, 1996). This discrepancy could be explained by time taken to translate and
fold GFP so that it can be bound by an antibody for recognition. Nonetheless, in later embryos
treated with control RNAi, we see EGA-GFP signal in the somatic cells but not in P4 (Figure
3.2). On the other hand, treatment with top-2/capg-2 double RNAi resulted in the detection
of EGA-GFP in P4 as well as the somatic cells of the embryos (Figure 3.2). This provides
additional evidence for the derepression of transcription in the P-lineage when TOP-2 and
condensin II are lost.

Aberrant expression of EMS specific genes is observed in the P-lineage when TOP-2
and condensin II are depleted.
Data presented thus far demonstrate aberrant transcriptional activity in P2, P3, and
P4 after depletion of TOP-2 or CAPG-2. To obtain additional evidence for the misregulation
of gene expression in these cells, we performed the in-situ hybridization chain reaction
(HCR), also known as RNA-FISH (Choi et al, 2016), to detect and localize mRNAs in early
embryos. For this, we focused on 4-cell embryos, and we started by looking at the expression
pattern of the somatic gene vet-6, which has previously been shown to be expressed
exclusively in EMS at the 4-cell stage (Seydoux et al., 1996). In wild-type embryos, vet-
6 expression was indeed localized to the EMS blastomere, with 100% of samples showing
vet-6 transcript signals in EMS. On the other hand, only 7.5% of wild-type embryos had vet-
6 signal in P2. Upon treatment with top-2 RNAi, however, most of the samples (57.5%)
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had vet-6 transcripts in P2 (Figure 3.3A). We also examined the expression pattern of an
additional transcript, F58E6.6. Similar to vet-6, F58E6.6 transcripts are seen predominantly
in EMS in wild-type embryos, with only 10-20% of our wild-type samples showing transcript
signals in ABa, ABp, or P2. After TOP-2 or CAPG-2 depletion, however, 50% and 82%,
respectively, of the embryos examined were now expressing F58E6.6 in P2(Figure 3.3B and
S3.2A&B). As a positive control, we depleted PIE-1 and found that both vet-6 and F58E6.6
transcripts were expressed in P2 in 85% and 100% of our samples (Figures 3.3A and
S3.2B&C). We conclude that depletion of TOP-2 and condensin II results in the misexpression
of EMS genes in the P-lineage of early embryos.

It has been reported that in pie-1 mutants, P2 loses its germline fate and its
descendants adopt the fate of EMS due to transcriptional reprogramming in P2 (Mello et al,
1992). With us reporting the misexpression of EMS specific genes in P2 in samples depleted
of TOP-2 and condensin II, we then asked if this depletion resulted in a similar EMS fate
adoption by P2. To test this, we utilized an EMS fate reporter strain where GFP driven by the
end-3 promoter marks EMS descendants with endoderm fates in C. elegans embryos (Maduro
et al, 2005) but were unable to detect any fate switch (data not shown). While this suggests
that fate change from P2 to EMS is unlikely to occur when TOP-2 and condensin II are lost,
more in-depth investigation into the fates of the P-lineage after TOP-2 and condensin II is
needed to make formal conclusion about the fates of these cells. Such works are currently
underway in our laboratory.

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The H3K9me pathway is dispensable for genome silencing in the P-lineage.
Data thus far have focused on the role of TOP-2/condensin II in transcriptional
control within the P-lineage. For the GCC pathway, TOP-2 and condensin II collaborate with
the H3K9me pathway to repress transcription, and thus we next explored the possibility that
H3K9me pathway components are operational during P-lineage silencing. One characteristic
of GCC-mediated silencing is the massive accumulation of H3K9me3 marks on chromatin,
and this is observed in Z2/Z3 of late embryos, in starved L1s, and in proximal oocytes (Belew
et al, 2021; Belew and Chien et al, 2023). To determine if H3K9me marks also accumulate in
transcriptionally silenced cells of the P-lineage, we stained early embryos for H3K9me3.
Figure 3.4A shows representative images for both early and late zygotes. Importantly, we
have previously shown that the chromatin of pre-fertilization oocytes is heavily decorated
with H3K9me3 marks, such that the immunofluorescence signal for H3K9me3 overlaps
entirely with the DNA-based DAPI signal (Belew and Chien et al, 2023). This is not the case
for nuclei in one-cell embryos, where H3K9me3 signal is present, but is limited to small
patches scattered along the chromosomes. By contrast, H3K9me3 signals are much stronger
on polar bodies, where they overlap with much of the DAPI signal (Figure 3.4A). These data
suggest that, upon fertilization, H3K9me3 marks are rapidly erased on much of the
chromatin inside the female pronucleus, but not on the chromatin in polar bodies. We went
on to examine 4- and 8-cell embryos, comparing P2 and P3 to their sister cells EMS and C. All
four cell types showed a similar pattern of H3K9me3 foci scattered throughout the
chromatin, and quantification of these signals showed that P cells did not accumulate
H3K9me3 marks to a greater extent than their transcriptionally active sisters (Figure 3.4B).
Consistent with a lack of massive accumulation of H3K9me3 marks in the P-lineage we also
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observed that set-25 mutant embryos could readily silence transcription in the P-lineage, as
assessed by RNAPIIpSer2 staining (Figure 3.4C). In both oocytes and starved L1s the SET-25
methyltransferase plays a crucial role in genome silencing (Belew et al, 2021, Belew and
Chien et al, 2023). Based on these data, we conclude that GCC pathway per se is not required
for genome silencing in the P-lineage, and thus that TOP-2 and condensin II mediate silencing
in a manner independent of heterochromatin assembly.

PIE-1 and TOP-2 work in parallel to repress transcription in the P-lineage.
So far, we and others have shown that 3 separate factors repress transcription in the
P-lineage: PIE-1, TOP-2 and condensin II. In a final stretch of experiments, we asked if TOP-
2 and condensin II work together with PIE-1 or if they work in parallel to repress
transcription. To address that, we wished to aberrantly express PIE-1 in the somatic cells
(Aba, ABp and EMS) of 4-cell embryos to repress transcription in those cells and ask if TOP-
2 and condensin II are required for this PIE-1 dependent transcriptional silencing. Previous
work has shown that the exclusive localization of PIE-1 in the P-lineage of early embryos is
achieved in part by its degradation from the somatic cells. The E3 ubiquitin ligase subunit
ZIF-1 is expressed exclusively in somatic cells and is required for PIE-1 degradation. MEX-5
and MEX-6 promote ZIF-1 expression in somatic cells by antagonizing its repressors MEX-3,
POS-1, and SPN-4 all of which actively repress ZIF-1 expression in the P-lineage (Figure
S3.3A; Oldenbroek et al, 2012). Therefore, we reasoned that the depletion of MEX-5/6 using
RNAi should result in ZIF-1 repression and, consequently, abolish the restriction of PIE-1 to
P-lineage cells of early embryos. That is what we saw when we used a strain that expresses
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PIE-1::GFP to track PIE-1 localization as after depleting MEX-5/6. In mex-5/6 RNAi samples
PIE-1 is expressed in all cells of 4-cell embryos (Figure S3.3B). This observation agrees with
previously documented pattern of PIE-1 expression when MEX-5/6 are lost (Schubert et al,
2000). We also observed that in mex-5/6 RNAi treated embryos, P-granules are present in
all cells which indicates that besides PIE-1 other germline proteins also get evenly
distributed across the embryo (Figure S3.3B). This means that cells in a mex-5/6 RNAi
embryo are of mixed somatic and germline fate which we make note of in our quantification
of Figures S3.3B and 3.5. Next, we wanted to see if the uniform distribution of PIE-1 across
all cells resulted in the loss of RNAPIIpSer2 marks in those cells. It has been previously
reported that expression of PIE-1 is sufficient to repress transcription in otherwise
transcribing cells (Seydoux et al, 1996; Batchelder et al, 1999). Consistent with this, mex-5/6
RNAi embryos had lost all RNAPIIpSer2 signal from all of their cells (Figure S3.3B).

We then wanted to see if TOP-2 and condensin II were required for this PIE-1
mediated silencing. Unfortunately, additional RNAi treatments, on top of mex-5/6 RNAi, were
not giving us the phenotypes that are normally associated with the treatments. To get around
this issue, we used a temperature sensitive mutant strain for top-2 (top-2(it7); Jaramilo-
Lambert et al, 2016) as it enabled us to attenuate TOP-2 without the use of RNAi. As shown
in Figure 3.5 control RNAi panels, shifting the top-2 TS mutans to a non-permissive
temperature of 24
o
C resulted in loss of TOP-2 function and aberrant transcription in P2
which is consistent with the data presented in Figure 3.1. When mex-5/6 RNAi animals were
shifted to 24
o
C, the inactivation of TOP-2 did not reverse the transcriptional repression by
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PIE-1 in any of the cells (Figure 3.5 - mex-5/6 RNAi panels). From this, we conclude that PIE-
1 does not work with TOP-2 in a linear fashion to repress transcription in early C. elegans
embryos.

An interesting observation is the fact that TOP-2 represses transcription exclusively
in germline precursor cells. In mixed fate cells in mex-5/6 RNAi embryos, the inactivation of
TOP-2 by shifting TS mutants to nonpermissive temperatures doesn’t result in aberrant
RNAPIIpSer2 signal in any of the cells. On the contrary, in control RNAi samples, TOP-2
inactivation leads to RNAPIIpSer2 signal accumulation in P2 (Figure 3.5). This shows that
proper inheritance of the germline fate seems to be required for TOP-2 to function as a
transcriptional repressor. This may be due to the presence of germline-specific TOP-2 co-
factors that enable it to repress transcription or soma-specific inhibitors of TOP-2. Testing
this hypothesis and identifying potential activators/repressors is an exciting avenue our
group is currently pursuing.

All in all, in this study we asked if the GCC pathway, known to silence the genomes
PGCs of starved L1s, also silences gene expression in P-lineage cells during germline
specification. The answer was clear, that GCC per se is not involved, as some of the hallmarks
of GCC events – for instance, the massive accumulation of H3K9me marks, a requirement for
SET-25 in silencing, and most notably extreme chromatin compaction – are not occurring in
P cells. We did, however, reveal a role for TOP-2/condensin II in genome silencing during
germline specification. It thus appears that TOP-2/condensin II can work independently of
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the H3K9me pathway in regulating transcription on a global level. Previous work had
identified PIE-1 as required for genome silencing in P2, P3, and P4, and thus TOP-2 and
condensin II are just the second and third factors to be identified that perform this crucial
function. Finally, we also show that TOP-2 and PIE-1 work in parallel to repress transcription
in the P-lineage.

Perhaps the most obvious future direction is identifying the mechanisms for the
transcriptional repression by TOP-2 and condensin II. Works from our group have linked
both TOP-2 and condensin II to chromatin based transcriptional repression (Belew et al,
2021; Belew and Chien et al, 2023). Although there is no obvious large-scale chromatin
compaction in the P-lineage of wild-type embryos (Belew et al, 2021), nanoscale compaction
(compaction that cannot be distinguished by confocal microscopy due to the diffraction limit
of 250nm in resolution) cannot be ruled out. Moreover, a previous study, using FRET-based
microscopy, has shown that condensin II plays a role in facilitating nanoscale compaction in
adult C. elegans germline (Llères et al, 2017). Therefore, it is possible that TOP-2 and
condensin II repress transcription in the P-lineage by facilitating nanoscale compaction of
the genome in those cells. More work needs to be done to test this and other possibilities and
characterize the mechanism of action for TOP-2 and condensin II mediated silencing.

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Materials and Methods
C. elegans strains
N2 (wild-type), WMM1 ([pie-1::gfp::pgl-1 + unc-119(+)]; [(pAA64)pie-
1p::mCherry::his-58 + unc-119(+)] IV), WMM2 (ltls37 [(pAA64) pie-1p::mCherry::his-58
+unc-119(+)] IV; unc-4(e120) top-2(it7ts) II), MT17463 (set-25(n5021) III), WM330 (pie-
1(ne4301[pie-1::GFP]) III) and 1284 (rrrSi199[Pvet-4::NLS:gfp:gfp::vet-4 3’UTR; unc-
119(+)] II; rrrSi198[Pvet-4::NLS:gfp:gfp::vet-4 3’UTR; unc-119(+)] IV) strains were used in
this study. Worms were maintained on 60mm plates containing nematode growth media
(NGM) seeded with the E. coli strain OP50 or HT115. Worms were grown at 20
o
C and
propagated through egg preparation (bleaching) every 72 hours.

Bacterial strains
OP50 bacteria served as the primary food source. It was grown in LB media
containing 100 ug/ml streptomycin by shaking at 37
o
C overnight. 500 ul of the culture was
seeded on Petri-dishes containing NGM + streptomycin. HT115 bacteria grown in LB media
containing 100 ug/ml carbenicillin and 12.5 ug/ml tetracycline and seeded on NGM +
carbenicillin + tetracycline plates were also used as a source of food. Our RNAi strains were
obtained from the Ahringer library and verified by Sanger sequencing. Bacteria containing
dsRNA were streaked on LB-agar plates containing 100 ug/ml carbenicillin and 12.5 ug/ml
tetracycline and incubated at 37
o
C overnight. Single colonies were then picked and grown in
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25ml LB cultures with 100 ug/ml carbenicillin and 12.5 ug/ml tetracycline. 500 ul of this
culture was seeded on 60 mm Petri-dishes containing 5mM IPTG.

Egg preparation
Bleach solution containing 3.675 ml H2O, 1.2 NaOCl, and 0.125 ml 10N NaOH was
prepared. Adult worms were washed from plates with 5 ml of M9 minimal medium (22mM
KH2PO4, 22mM Na2HPO4, 85mM NaCl, and 2mM MgSO4). Worms were centrifuged at 1.9
KRPM for 1 minute and the excess medium was removed, then the bleach solution was
added. Eggs were extracted by vortexing for 30 seconds and shaking for 1 minute. This was
done a total of 3 times and worms were vortexed one last time. Then the eggs were spun
down at 1900 rpm for 1 minute and excess bleach solution was removed and the eggs were
washed 3 times with M9 minimal medium.

RNAi treatment
RNAi containing NGM plates were prepared as described in the “Bacterial strains”
section. For double RNAi treatments, RNAi cultures were mixed at a 1:1 ratio by volume.
HT115 cells transformed with an empty pL4440 vector was used as a negative control. RNAi
conditions used in this study and tests for their efficacy is described below:

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top-2 RNAi
L1 worms were plated on HT115 food plates for the first 24 hours and were then moved to
plates seeded with top-2 RNAi for the remaining 48 hours. Embryonic lethality was observed
at >90%.
capg-2 RNAi
Worms were grown on HT115 food plates for the first 24 hours and were moved to plates
containing capg-2 RNAi for the remaining 48 hours. An embryonic lethality of 80%-100%
was seen with this RNAi treatment.
top-2/capg-2 double RNAi
Worms were grown on HT115 food plates for the first 24 hours and were transferred to top-
2/capg-2 double RNAi plates for the next 48 hours. Embryonic lethality ranged from 90%-
100% for this RNAi treatment.
set-25 RNAi
Worms were grown on plates containing set-25 RNAi for the entirety of their life cycle. Some
of the adult worms were bleached and an L1 chromatin compaction assay was performed on
the resulting larvae to test RNAi efficacy. Please see Belew et al., 2021, for details on the L1
compaction assay.
pie-1 RNAi
Worms were grown on plates containing pie-1 RNAi for the entirety of their life cycle. An
embryonic lethality of 100% was observed for this RNAi.
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mex-5/6 RNAi
Worms were grown on HT115 food plates for the first 48 hours of their life and were then
switched to mex-5/6 RNAi plates for the remaining 24 hours. An embryonic lethality of
100% was observed for this RNAi treatment.

Antibodies and dilutions
RNAPIIpSer2: Rabbit antibody from Abcam (ab5095, Cambridge, Massachusetts) was
used at a dilution of 1:100. GFP: mouse Mab #3580, from EMD Millipore, was used at 1:500.
H3K9me3: Rabbit antibody from Abcam (ab176916, Cambridge, Massachusetts) was used at
a dilution of 1:1000. P-granules: mouse Mab K76, from the Developmental Studies
Hybridoma Bank, was used neat. Secondary antibodies: Alexa Fluor conjugated secondary
antibodies from Invitrogen (Thermofisher Scientific, Waltham, Massachusetts) were used at
a dilution of 1:200.

Immunofluorescence staining
Adult worms were first washed off plates with 10 ml of M9 minimal medium and
washed 3 more times. Then, they were centrifuged at 1.9 KRPM and the excess medium was
removed. 20 ul of media containing about 50 worms were spotted on a coverslip and 3 ul of
anesthetic (20mM Sodium Azide and 0.8M Tetramisole hydrochloride) was added to
immobilize them. Worms were dissected using 25Gx5/8 needles (Sigma Aldrich, St. Louis,
Missouri). To release early embryos, worms were cut once midway through their length. The
144

coverslip was then mounted onto poly-L-lysine covered slides and let rest for 5 minutes.
Slides were put on dry ice for 30 minutes. Samples were then freeze-cracked by flicking the
coverslips off for permeabilization.

For RNAPIIpSer2 staining experiments, once samples are permeabilized, slides were
put in cold 100% methanol for 2 minutes and then fixing solution (0.08M HEPES pH 6.9,
1.6mM MgSO4, 0.8mM EGTA, 3.7% formaldehyde, 1X phosphate-buffered saline) for another
30 minutes. After fixing, slides were washed three times with TBS-T (TBS with 0.1% Tween-
20) and were blocked for 30 minutes with TNB (containing 100mM Tris-HCl, 200 mM NaCl,
and 1% BSA). Primary antibodies were then applied at the dilutions described above in TNB
and slides were incubated at 4
o
C overnight.

For H3K9me3 and GFP staining experiments, permeabilized samples were put in cold
100% methanol for 10 seconds and then fixing solution (0.08M HEPES pH 6.9, 1.6mM
MgSO4, 0.8mM EGTA, 3.7% formaldehyde, 1X phosphate-buffered saline) for 10 minutes.
After fixing, slides were washed three times with TBS-T (TBS with 0.1% Tween-20) and were
blocked for 2 hours with TNB (containing 100mM Tris-HCl, 200 mM NaCl, and 1% BSA)
supplemented with 10% goat serum. Primary antibodies were then applied at the dilutions
described above in TNB and slides were incubated at 4
o
C overnight.

145

On the next day, the slides were washed 3 times with TBS and slides were incubated
with secondary antibodies and Hoechst 33342 dye for 2 hours at room temperature. Slides
were washed 3 times with TBS, mounting medium (50 % glycerol in PBS), and coverslips
were applied and sealed with Cytoseal XYL (Thermofisher).

HCR (In Situ Hybridization Chain Reaction)
A kit containing a DNA probe set, DNA hybridized chain reaction (HCR) amplifier
hairpins, and hybridization, wash, and amplification buffers were purchased from Molecular
Instruments (molecularinstruments.com). Genes that were examined were vet-6 and
F58E6.6. DNA was visualized with Hoechst 33342 dye. Embryos were prepared by bleaching
and were immediately spotted on poly-L-lysine coated slides. Coverslips were applied and
slides were freeze-cracked to permeabilize samples. Immediately after freeze-cracking, 500
ul of 100% ice-cold methanol was applied over the samples. Slides were dried off by tilting
slides, 1 ml of 4% paraformaldehyde (PFA) was added and samples were incubated in a
humidity chamber for 10 minutes. Samples were then washed 3 times with 100 ul of PBS-T.
A 1:1 solution of probe hybridization buffer (PHB) and PBS-T was added to the samples, and
they were incubated for 5 minutes at room temperature. Samples were then prehybridized
with PHB for 30 minutes at 37oC and DNA probes (at a final concentration of 2 picomoles
per 500 ul of PHB) were added to the samples and were incubated overnight at 37oC.

The next day, samples were washed 4 times with probe wash buffer (PWB) at 37oC
with 15 minutes of incubation for each wash. They were then washed three more times with
146

5xSSCT at room temperature. Samples were pre-amplified with Amplification buffer for 30
minutes at room temperature. Probe amplifier hairpins were snap cooled by heating to 95oC
for 90 seconds and putting in a dark drawer for 30 mins then were added to the sample.
Worms were incubated with the hairpins overnight in a dark drawer.

On the third day, samples were washed with 5xSSCT and incubated with Hoechst-
33342 (1:5000 dilution) for 15 minutes. Finally, samples were mounted on poly-L-lysine
coated slides and imaged.

Immunofluorescent imaging
All slides were imaged using an Olympus Fluoview FV1000 confocal microscope using
Fluoview Viewer software. A magnification of 600x (60x objective and 10x eyepiece
magnifications) was used. Laser intensity was controlled for experiments to achieve
consistency among samples.

147

Figures and Figure Legends

148

Figure 3.1: Loss of TOP-2 or CAPG-2 allows for active RNAPII in the P-lineage of early
embryos.
Early embryos from N2 worms treated with control, top-2, capg-2 or pie-1 RNAi were fixed
and stained for P-granules (green), RNAPIIpSer2 (red) and DNA (blue). Shown on top are the
entire embryos and the P-cell is magnified below. Quantification of the data is shown at right.
149

150

Figure 3.2: EGA-GFP gets misexpressed in P4 when TOP-2 and CAPG-2 are depleted.
Early embryos from 1284 worms were treated with either control or top-2/capg-2 RNAi.
Samples were fixed and stained for P-granules (white), EGA-GFP (green) and DNA (blue). 20
samples were analyzed over 2 independently performed replicates. EGA-GFP signal was
aberrantly present in P4 of embryos treated with top-2/capg-2 RNAi. Quantification of the
data is shown below.

151

152

Figure 3.3: EMS specific genes are aberrantly expressed in the P2 cell of 4-cell embryos
after loss of TOP-2 or CAPG-2.
(A) 4-cell embryos from N2 animals treated with control, top-2 or pie-1 RNAi were fixed
and HCR was performed to probe for vet-6 mRNA (white). DNA was stained using
Hoechst-33342 (blue). Representative images are shown and percentage of samples
with vet-6 transcripts in P2 are shown below (n=40).
(B) 4-cell embryos from N2 animals treated with control, top-2 or capg-2 RNAi were fixed
and HCR was performed to probe for F58E6.6 transcripts (white). DNA was visualized
using Hoechst-33342 dye (blue). P2 nuclei are outlined with red dashed line while
somatic nuclei are outlined by white dashed line. Representative images are shown
and quantification of F58E6.6 signal in each blastomere is shown on the right.

153

154

Figure 3.4: The H3K9me heterochromatin pathway is not required for transcriptional
repression in the P lineage.
(A) The indicated zygotes from N2 worms were fixed and stained for H3K9me3 (red) and
DNA (blue). Shown below the entire embryos are magnified views of the pronuclei
and polar bodies (PB). Note that the H3K9me3 signal overlaps with the Hoechst signal
more extensively on the polar bodies than within the pronuclei.
(B) Same as (A) except two- and four-cell embryos were examined. Quantification of
H3K9me3 foci for P2 and P3 (germ) or EMS and C (soma) is shown below.
(C) Four-cell embryos, either wild type (N2) or set-25 mutants (strain MT17463), were
fixed and stained for RNAPIIpSer2 (red) and P-granules (blue). 20 samples were
analyzed over 2 independent experiments. Both wild-type and set-25 mutants had no
H3K9me3 signal in P2.

155

156

Figure 3.5: PIE-1 does not require TOP-2 to repress transcription in C. elegans early
embryos.
4-cell embryos from WMM2 (top-2 TS) animals treated with either control or mex-5/6 RNAi
were optionally shifted to the nonpermissive temperature of 24oC for 24 hours or left at the
permissive 15oC. Then samples were fixed and stained for P-granules (white), RNAPIIpSer2
(red) and DNA (blue). 20 samples were analyzed over 2 independent replicates. Attenuation
of TOP-2 via temperature shift does not reverse the transcriptional silencing by PIE-1 in mex-
5/6 RNAi embryos. Quantification of the data is presented below.

157

Supplementary Figures

158

Figure S3.1: RNAPII-mediated transcription is still repressed in one- and two-cell
embryos after loss of TOP-2 or SET-25.
One- and two-cell embryos from N2 worms treated with the indicated RNAi were fixed and
stained for P-granules (white), RNAPIIpSer2 (red) and DNA (blue). No active RNAPII was
observed in the more than 20 samples we examined for both one- and two-cell embryos over
2 replicates. Note the RNAPIIpSer2 signals in the adjacent, older embryo in the upper right
panel – this shows that the staining worked efficiently.

159

160

Figure S3.2: Control experiments for somatic gene expression in 4-cell embryos.
(A) Samples were prepared in the same way as Figure 3.3B except RNAi against pie-1 was
utilized.
(B) The same panels as those shown in Figure 3.3B are displayed without the dashed
circles.
(C) A summary of the data shown in Figures 3.3B and S3.2A.

161

162

Figure S3.3: Depletion of MEX-5 and MEX-6 results in PIE-1 presence in all blastomeres
of early embryos.
(A) Schematic showing the control of PIE-1 asymmetry in early embryos by MEX-5 and
MEX-6. See results section for details.
(B) 4-cell embryos from WM330 animals optionally treated with either control or mex-
5/6 RNAi were fixed and stained for P-granules (white), PIE-1::GFP (green),
RNAPIIpSer2 (red) and DNA (blue). Images of the same embryo stained for PIE-
1::GFP and RNAPIIpSer2 are shown side by side. 20 samples were analyzed over 2
independent replicates. Depletion of mex-5/6 results in the loss of asymmetric
distribution of PIE-1 to the germline. PIE-1 is present in all blastomeres and that is
sufficient to repress transcription in the whole embryo as shown by low RNAPIIpSer2
levels.

163

Contributions
Below is a table of the figures where I contributed to by collecting/analyzing the data and
making figures.
CHAPTER 1 CHAPTER 2 CHAPTER 3
Figure 1.1A, C, D Figure 2.1 B, C Figure 3.1 (capg-2 pannel)
Figure 1.2 A-F Figure 2.2 B Figure 3.2
Figure 1.3 A, B, C, E, F Figure 2.3 A, B Figure 3.4
Figure 1.4 A-C Figure 2.4 A, B Figure 3.5
Figure 1.5 A-C Figure 2.5 A, C Figure S3.1
Figure S1.2 A-D Figure 2.6 A (set-25 pannel), B FigureS3.3 B
Figure S1.3 B Figure 2.7
Figure S1.4 Figure 2.8 A-C
Table S1.1 Figure S2.1 A, B
Figure S2.2 B
Figure S2.3 B, C
Figure S2.4 A, B
Figure S2.5

164

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Asset Metadata

Creator Belew, Mezmur Dawit (author)

Core Title Identification of factors that underly genome-wide transcriptional repression during the development of the Caenorhabditis elegans germline.

Contributor Electronically uploaded by the author (provenance)

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Degree Conferral Date 2023-08

Publication Date 06/29/2023

Defense Date 06/20/2023

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag adult gonads,C. elegans,cell biology,condensin II,genome-wide transcriptional repression,germline precursor cells,global chromatin compaction (GCC) pathway,heterochromatin,L1 starvation,L1s,maturing oocytes,Molecular Biology,OAI-PMH Harvest,oocytes,PIE-1,primordial germ cells,topoisomerase 2,Transcription,transcriptional regulation

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Language English

Advisor Michael, W. Matthew (committee chair), Calabrese, Peter (committee member), Forsburg, Susan (committee member), Philips, Carolyn (committee member)

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Abstract (if available)

Abstract The proper transmission of genetic material from generation to generation is an important aspect of life. In the case of multicellular animals that reproduce sexually, specialized cell types known as germ cells, are tasked with this. To ensure the fidelity of the message transmission, germ cells are under constant surveillance for aberrances and employ multiple mechanisms to avoid them. One mechanism that is well conserved among species is the tight regulation of transcription in the germ cells during the development of the organism. To avoid the expression of unwanted genes that may alter the germline fate of these cells, animals employ gene silencing at specific targets or at a genome-wide level.

In the nematode Caenorhabditis elegans (C. elegans), the development of the germline is characterized by cycles of genome-wide transcriptional activation and repression. In early embryos, the P-lineage of cells, that gives rise to the germline, is kept transcriptionally quiescent by PIE-1. Another round of global transcriptional repression is observed in primordial germ cells (PGCs) when a newly hatched stage 1 larva (L1) is starved. Unlike the repression in early embryos, this relies on changes in the chromatin landscape rather than a single factor facilitating it. Loss of active chromatin marks such as H3K4me, H4K8ac, and H4K16ac have been reported. Yet, what underlies this change in chromatin architecture and the extent of the change in chromatin shape still require characterization.

In this study, we delve into the chromatin-based transcriptional repression in starving L1s. We show that this repression is accompanied by a physical compaction of the PGC chromatin where the DNA is compacted into bundles that are located at the nuclear periphery. We also show that this compaction is driven by a novel pathway that entails a partnership between topoisomerase II (TOP-2), the condensin II complex, and various heterochromatin-related proteins - such as the H3K9 methyltransferases MET-2 and SET-25 and the H3K9me mark reader proteins CEC-4 and HPL-2. We call this pathway the global chromatin compaction (GCC) pathway. This pathway is triggered during L1 starvation by the low-energy sensing proteins AAK-1 and AAK-2 which are homologs of the mammalian AMPK. Next, we show that the components of the GCC pathway are also required to repress transcription in maturing proximal oocytes in the adult germline. We also report that the known transcriptional repressor, PIE-1, also functions in maturing oocytes in addition to P-lineage cells of early embryos. Finally, we show that the TOP-2/condensin II axis works in tandem with the PIE-1 to repress transcription in the P-lineage cells of early embryos. All in all, these studies provide insights into the factors that underly genome-wide transcriptional silencing at different points during the development of C. elegans germline.