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The organization of small RNA pathways within C. elegans germ granules: mutator foci formation, regulation, and interaction

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The organization of small RNA pathways within C. elegans germ granules: mutator foci formation, regulation, and interaction

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Content Copyright 2021 Celja Jae Churches Uebel

THE ORGANIZATION OF SMALL RNA PATHWAYS
WITHIN C. ELEGANS GERM GRANULES

Mutator foci formation, regulation, and interaction

by

Celja Jae Churches Uebel

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

December 2021
ii
ACKNOWLEDGEMENTS

Since the age of 12, I have wanted to be a scientist. The journey to and through
graduate school has been a long time in the making, but would not have been possible
without the strong community of family, friends, and mentors that have ballasted me on
the way. First and foremost, my utmost gratitude goes to my doctoral advisor, Dr. Carolyn
Phillips, for her mentorship, support, and encouragement throughout my PhD. Carolyn’s
mentorship helped me to thrive as a researcher by providing the perfect combination of
early intellectual guidance and subsequent room to grow in following my own scientific
curiosities. Thank you for teaching me the art of storytelling through scientific figures and
for allowing me to be a well-rounded person outside of academia, creating a positive and
balanced graduate school experience. However, I would not be in graduate school without
my undergraduate advisor, Dr. Lisa Wrischnik, who remains my inspiration in pursuing
professorship. Wrisch was witness to my first scientific successes in lab, and importantly,
my first failures. She taught me tenacity, creativity, and critical thinking to overcome the
challenges of lab, and I am honored to carry on her quirk of pipetting with the index finger
rather than the thumb. I hold a deep, deep gratitude for both my advisors’ mentorship.
I also hold a sincere appreciation for the members of my committee. To Dr. Fabien
Pinaud, thank you for your willingness to collaborate on 3D-STORM imaging, which
ultimately spurred me to pursue other high-resolution imaging methods and furthered my
love of microscopy. To Dr. Sean Curran, thank you for bringing the nematode perspective
in your many thoughtful questions and suggestions for my research. And finally, to Dr.
Matt Pratt, whose Advanced Chemical Biology course reminded me to view biological
iii
processes from multiple scientific perspectives. Thank you all for the great advice and
encouragement throughout my PhD.
A huge thanks is due to my peers at USC. In the Phillips lab: Thank you, Kevin, for
your grandfatherly and steadfast advice. Alicia, I am constantly looking up to you for
encouragement that I, too, can become a professor one day. Thank you for helping me
navigate postdoc applications and for always speaking your mind. Kat, thank you for
being the backbone of the lab and for being a genuinely kind person. Dylan, thank you
for matching my energy in lab, both with jokes and with rigorous scientific discussion. To
the Phillips lab as a whole: thank you for the camaraderie, laughs, Cheese Fridays,
encouragement, and friendship. I wish our newest members luck on their own journeys! I
also want to recognize my cohort friends Justin, Katie, Calista, Gary, James and Anupam
for the support and friendship outside of the lab. Thank you also to my friends in the
molecular biology program as a whole, to whom a single line of appreciation is not nearly
enough to express my gratitude!
Within the department, I want to thank the administrators and staff whose job
enables us to focus on our research. Thank you, Paloma, Doug, Katie, Jen, Rokas, and
others. A special thanks to Miriam, for gracing RRI with her warmth, smile, and cheerfully
whistled tune.
I would also like to bring attention to the innate privilege in my life which has
opened doors that may be unfairly closed to others. In part, my privilege manifests in my
ability to apply for and receive national scholarships as a US citizen, in being able to
volunteer unpaid hours in an undergraduate lab to gain research experience, and in
having peers and mentors that look like me. I want to recognize this privilege partly to
iv
highlight the inequity on which systems of academia are currently built and partly to
remind myself and the department to continue to push for accountability and equity in
science.
Lastly, words are not sufficient to fully express my gratitude to my family. To my
parents, Laurie and Chris Uebel, who have always empowered me to pursue my own
passions. Thank you for teaching me that hard work and determination has the ability to
outlast innate talent. To my brother, Nicholas Uebel, who inherited most of the innate
talent. Thank you for making me laugh and inspiring me with your empathy, intellect, and
imagination. To my extended family, who are my support system and my people. Finally,
to my husband, Dr. Nathan Daniel Churches, who continually inspires me to be a better
mentor, scientist, and person. Thank you for your unconditional encouragement and love.
v
TABLE OF CONTENTS
ACKNOWLEDGEMENTS ii
LIST OF FIGURES vii
ABSTRACT ix
INTRODUCTION 1
BRIEF OVERVIEW 1
SMALL INTERFERING RNA PATHWAYS 4
SMALL RNA PATHWAY ORGANIZATION 19
CREATING A SPATIAL MODEL OF RNA SILENCING 33
CHAPTER 1: MUTATOR FOCI FORMATION 35
PREFACE 35
ABSTRACT 35
INTRODUCTION 36
RESULTS 39
DISCUSSION 50
METHODS 55
ACKNOWLEDGMENTS 62
CHAPTER 1 FIGURES 63
MESO-CHAPTER 1 77
PREFACE 77
ABSTRACT 77
RESULTS 78
METHODS 80
MESO-CHAPTER 1 FIGURE 83
CHAPTER 2: MUTATOR FOCI REGULATION 84
PREFACE 84
ABSTRACT 84
INTRODUCTION 85
RESULTS 88
DISCUSSION 96
MATERIALS AND METHODS 100
ACKNOWLEDGEMENTS 103
CHAPTER 2 FIGURES 105
MESO-CHAPTER 2 113
PREFACE 113
ABSTRACT 113
RESULTS 114
MATERIALS AND METHODS 116
MESO-CHAPTER 2 FIGURE 117
vi
CHAPTER 3: NUAGE INTERACTION AND ORGANIZATION 118
PREFACE 118
ABSTRACT 119
INTRODUCTION 119
RESULTS 125
DISCUSSION 133
MATERIALS AND METHODS 136
ACKNOWLEDGEMENTS 143
CHAPTER 3 FIGURES 144
SUPPLEMENTAL CHAPTER 1 154
RESULTS 154
MATERIALS AND METHODS 156
SUPPLEMENTAL CHAPTER 1 FIGURES 160
SUPPLEMENTAL CHAPTER 2 162
RESULTS 162
MATERIALS AND METHODS 165
ACKNOWLEDGEMENTS 167
SUPPLEMENTAL CHAPTER 2 FIGURES 168
SUPPLEMENTAL CHAPTER 3 170
RESULTS 170
MATERIALS AND METHODS 172
SUPPLEMENTAL CHAPTER 3 FIGURES 174
REFERENCES 176

vii
LIST OF FIGURES

INTRODUCTION
Figure 1. The C. elegans small interfering RNA pathway 6

CHAPTER 1
Figure 1. MUT-16 and orthologs contain a high degree of predicted disorder 63
Figure 2. Susceptibility of mut-16 deletion worms to somatic and germline
RNAi
64
Figure 3. The C-terminal region of MUT-16 is necessary for Mutator foci
formation
65
Figure 4. Distinct regions of MUT-16 recruit each of the other Mutator
proteins
66
Figure 5. The C-terminal region of MUT-16 is sufficient for foci formation 67
Figure 6. Concentration dependence of Mutator foci 68
Figure 7. Mutator foci have liquid-like properties 69
Figure S1. Distribution of Q, N, and P residues in MUT-16 orthologs 70
Figure S2. Requirements for RRF-1, NYN-1, and RDE-8 localization to
Mutator foci
71
Figure S3. Response of GFP- and mCherry-tagged strains to somatic and
germline RNAi
72
Figure S4. Distinct MUT-16 regions are required for the localization of the
other Mutator proteins
73
Figure S5. Regions B and C of MUT-16 are required for interaction with
MUT-2 in vitro
75
Figure S6. MUT-16 foci are temperature dependent 76

MESO-CHAPTER 1
Figure 1. PGL-1 fluorescently tagged with mKate2 or mTagBFP2 has distinct
phase-separation dynamics compared to PGL-1::GFP
83

CHAPTER 2
Figure 1. Distinct Mutator foci appear in the Z2/Z3 progenitor germ cells 105
Figure 2. Mutator foci are present in L1-L3 larval stages 106
Figure 3. Mutator foci are present throughout spermatogenesis and are
found in the cytoplasm of spermatids
107
Figure 4. Mutator foci integrity partially relies on RNA 108
Figure 5. Mutator foci intensity varies along the adult germline 109
Figure S1. Mutator foci are weakly present in the 30-cell stage 110
Figure S2. Mutator foci dissipate after ama-1 RNAi 111
viii
Figure S3. Variation in Mutator foci intensity across gonad regions 112

MESO-CHAPTER 2
Figure 1. SIMR foci are numerous and bright in the Z2/Z3 progenitor germ
cells
117

CHAPTER 3
Figure 1. Mutator foci and P granule separation is independent of nuclear
association and can to be reestablished after perturbation
144
Figure 2. P granules form unique pocket morphologies in the mid and late
pachytene
146
Figure 3. Nuage compartments exhibit a hierarchical stoichiometry 147
Figure 4. P granule pockets exhibit an exterior-to-interior organization 148
Figure 5. Exchange of biomolecules is facilitated within P granule pockets 149
Figure 6. Working model of P granule pocket organization of nuage 150
Figure S1. Ectopic interaction and dissolution of granules is consistent
across multiple fluorescent tags
151
Figure S2. P granule pocket morphology is visible in widefield live imaging
and in untagged P granules
152
Figure S3. Further analysis of nuage compartment populations 153

SUPPLEMENTAL CHAPTER 1
Figure 1. MUT-16 is posttranslationally modified 160

SUPPLEMENTAL CHAPTER 2
Figure 1. Distribution of MUT-16 protein is non-uniform 168
Figure 2. Mutator foci have complex morphologies 169

SUPPLEMENTAL CHAPTER 3
Figure 1. Visualization of endogenously silenced RNA 174
Figure 2. Visualization of oma-1 targeted by RNAi 175

ix
ABSTRACT

Small RNA pathways are critical regulators of gene expression in eukaryotic cells.
In the nematode C. elegans, small RNAs target and downregulate complementary RNA
to ensure proper gene expression and silence deleterious transcripts. Robust silencing is
achieved by small RNA amplification, which is dependent upon a perinuclear
membraneless compartment in germ cells called Mutator foci. Disruption of Mutator foci
causes chromosomal nondisjunction, temperature-sensitive sterility, and transposon
activation.
My doctoral work initially investigates the molecular formation of Mutator foci. First,
in examining different regions of MUT-16, a protein which nucleates Mutator foci, I
discover important functional subdivisions of the MUT-16 protein which are necessary for
protein-protein interaction and foci formation. I further probe the biophysical properties of
MUT-16 and discover that Mutator foci are liquid-like condensates which assemble by
phase separation. Mutator foci are adjacent to two additional phase-separated
condensates: P granules, which interact with newly exported RNA, and Z granules, which
are necessary for siRNA inheritance. Building on previous studies, my inaugural work
proposes that RNA silencing is facilitated by multiple phase-separated condensates at
the nuclear periphery of germ cells.
Next, I investigate the regulation of Mutator foci and define the spatiotemporal
appearance and localization of Mutator foci. I discover that Mutator foci arise in early
embryos, persist in all larval stages, and are present in spermatids, indicating a role in
paternal inheritance of small RNAs. I additionally demonstrate that RNA and the germline
x
cell cycle influence the morphology of Mutator foci and suggest that these factors may
influence the efficacy of RNA silencing in certain cellular environments.
Lastly, I probe the interaction of Mutator foci with adjacent perinuclear
compartments involved in small RNA pathways, which I refer to as C. elegans nuage. I
demonstrate that nuage interaction is independent of the germline environment and able
to be reestablished after disruption. I interrogate the spatial configuration of nuage
compartments with superresolution microscopy techniques and discover a previously
undescribed toroidal P granule morphology, in which some P granules surround Mutator
foci and other nuage compartments with a consistent exterior-to-interior organization.
Finally, I find that Mutator foci are the scarcest nuage compartment and that nuage
compartments exist in distinct populations, suggesting a functional subdivision of different
nuage assemblages. My final work aims to create a more accurate model of RNA
silencing through multiple phase-separated compartments. Together, my doctoral studies
begin to reveal how the complex and critical small RNA pathways are organized within C.
elegans nuage.
Copyright 2021 Celja Jae Churches Uebel
INTRODUCTION

BRIEF OVERVIEW
Regulatory RNAs silence transcripts via complex pathways
Small RNAs (sRNA) are tiny biomolecules with widespread regulatory
consequences. These anciently conserved regulatory elements are short, 18-30
nucleotide, non-coding stretches of ribonucleic acid (RNA) with the ability to silence
complementary transcripts. Targets of sRNA pathways include deleterious transcripts
such as viral RNA, selfish genetic elements, repetitive elements, and pseudogenes
(Claycomb 2014). Disruptions in sRNA pathways have dire consequences — in humans,
sRNA misregulation is linked to disease progression, drug resistance, and negative
prognosis of cancers (Zheng et al. 2010; Zhang et al. 2021b); in the fruit fly Drosophila,
sRNA mutants have shorter life spans, developmental defects, and fertility defects (Aravin
et al. 2001; Lim et al. 2011); and in the nematode Caenorhabditis elegans, the loss of
sRNA pathways results in temperature-sensitive sterility, genomic instability, and a
reduction in viable progeny (Collins et al. 1987; Smardon et al. 1999; Rogers and Phillips
2020b). Thus, small RNAs are a critical regulatory component of eukaryotic cells.
In filling such widespread regulatory roles, small RNA pathways have evolved
different mechanisms of silencing, multiple modes of biogenesis, and a suite of
specialized protein interactors to facilitate sRNA processing and transcript targeting.
Based on these characteristics, sRNAs have been broadly subdivided into three major
classes: micro-RNAs (miRNA), piwi-interacting RNAs (piRNA), and small interfering
RNAs (siRNA). In brief, the biogenesis of both miRNAs and siRNAs begins with double
stranded RNA (dsRNA) — via endogenously expressed stem-loop structures for miRNAs
2
(Ambros et al. 2003; Lee et al. 2004), or from viral dsRNA, hybridization of sense and
anti-sense transcripts, transposons, or RNA-dependent RNA polymerase activity for
siRNAs (Parrish et al. 2000; Watanabe et al. 2008; Vasale et al. 2010). In contrast,
piRNAs originate from single-stranded RNA precursors transcribed from discrete
genomic clusters (Aravin et al. 2006; Gainetdinov et al. 2018). For transcript silencing,
piRNAs and miRNAs target transcripts with partial complementarity, whereas siRNAs are
typically fully complementary but can tolerate a few mismatches (Montgomery et al. 2012;
Bagijn et al. 2012; Hutvanger and Zamore 2002). To add to the growing complexity, in
some organisms the initially produced “primary” siRNA will trigger the subsequent
production of a “secondary” siRNA species, resulting in further amplification of the
silencing signal (Aoki et al. 2007; Pak and Fire 2007; Vasale et al. 2010). Furthermore,
despite variation in nucleotide length, 5’ modification, and 3’ overhang, all three species
of small RNAs must be properly routed to and loaded onto a family of small RNA binding
proteins called Argonautes. This modular Argonaute-small RNA combination, known as
the RNA-induced Silencing Complex (RISC), then targets and downregulates the
transcripts as determined by the sequence of the loaded small RNA (Hutvanger and
Simard 2008). Finally, downregulation can occur both transcriptionally through deposition
of chromatin silencing marks, or post-transcriptionally through inhibition of translation,
recruitment of exonucleases, or direct Argonaute-mediated cleavage of the target RNA
(Elbashir et al. 2001; Gu et al. 2012). The currently known complexity of these pathways
is truly astounding.
Subcellular organization of complex pathways
On a molecular level, the complexity of these pathways is in direct conflict with the
fundamental challenge of subcellular organization. While the individual components of
3
small RNA pathways are well-characterized, we have a limited understanding of how
small RNA proteins and biomolecules are coordinated to facilitate efficient RNA silencing
within a crowded intracellular space.
The basis of my doctoral research aims to investigate the subcellular organization
of complex small RNA pathways. To this end, it was imperative to narrow the research
focus to a well-characterized pathway within a suitable model organism. An obvious
organismal selection was Caenorhabditis elegans, the long-studied and genetically
manipulable nematode in which the mechanism of small RNA-mediated RNA silencing
was first discovered (Fire et al. 1998). In addition to the ease of animal husbandry, the
self-fertilizing hermaphroditic worm enables straightforward production of genetic
homozygotes and many protocols have been established for CRISPR genome editing
(Brenner 1974; Dickinson and Goldstein 2016; Paix et al. 2017; Ghanta and Mello 2020).
Furthermore, in C. elegans, the small interfering RNA (siRNA) pathway stands out as
both particularly robust and remarkably complex. Around 26 putative Argonaute genes
have been identified, compared to the 5 Argonaute proteins in Drosophila and 8
Argonaute proteins in humans (Yigit et al. 2006; Hutvagner and Simard 2008).
Additionally, both exogenous and endogenous sources of dsRNA trigger primary sRNAs,
which can be further amplified into a heritable, secondary siRNA signal (Grishok et al.
2000; Pak and Fire 2007; Aoki et al. 2007; Vasale et al. 2010). Somehow, these multiple
unique Argonaute proteins properly associate with numerous distinct sRNA classes to
scan thousands of nascent transcripts and direct RNA silencing. For these characteristics,
I began to investigate the organization of the siRNA pathway within C. elegans to
understand how complex pathways efficiently function within a crowded cytoplasm.
4

SMALL INTERFERING RNA PATHWAYS
Early Discovery
The silencing effects of small interfering RNA pathways were described long
before the mechanism was formally understood. In the 1990s, two different research
groups attempted to overexpress purple pigment in petunias, and were instead met with
variegated or completely white flowers (van der Krol et al. 199; Napoli et al., 1990). At the
time, this phenomenon was known as “co-suppression” or post-translational gene
suppression. The molecular mechanism behind this silencing phenotype was elusive until
1998; when Andrew Fire and Craig Mello discovered that the injection of dsRNA could
potently silence C. elegans genes (Fire et al., 1998). This discovery, coined “RNA
interference” (RNAi), uncovered key ideas that laid the foundation for understanding
siRNA pathways: (1) the effectors of RNAi are produced from dsRNA specifically targeting
the mature mRNA and not genomic introns or promotors, (2) RNAi can be triggered by a
low concentration of injected dsRNA, indicating an amplification process, and (3) the
model organism C. elegans has a robust, precise, and heritable RNAi response. Quickly
after initial discovery in nematodes, RNAi was confirmed in Drosophila, Xenopus, plants,
and mice, demonstrating conservation of RNAi pathways across animal kingdoms
(Kennerdell and Carthew 1998; Waterhouse et al. 1998 Oelgeschlager et al. 2000;
Wianny and Zernicka-Goetz 2000). Mutations in C. elegans that disrupted RNAi, or de-
repressed key targets of RNAi, soon began to illuminate the molecular machinery of small
RNA pathways and establish C. elegans as a model organism with which to study these
regulatory RNAs (Tabara et al. 1999; Ketting et al. 1999). The following two decades
5
brought about characterization of RNAi protein components, pathway evolution,
regulatory mechanisms, and molecular technologies.

Classes of siRNAs
The culmination of more than twenty years research on RNAi has resulted in a
thorough characterization of the siRNA pathways. In C. elegans, the production of
“primary” small RNAs leads to a downstream amplification of “secondary” siRNAs (Pak
and Fire 2007; Aoki et al. 2007; Gu et al. 2009). Primary sRNAs can be endogenously-
derived siRNAs (endo-siRNAs) from protein coding genes, pseudogenes, repetitive
elements and duplications, or exogenously-derived (exo-siRNAs) from natural sources of
dsRNA, such as viral RNA, or artificially introduced dsRNAs (Fire et al. 1998; Lee et al.
2006; Fischer et al. 2013; Claycomb 2014). piRNAs can also act as primary sRNAs,
feeding into the downstream secondary siRNA pathways (Ruby et al. 2006; Das et al.
2008; Batista et al. 2008). Different classes of primary sRNAs and secondary siRNAs are
identified by the Argonaute with which they associate, the RNA length, and the 5’
sequence bias. Thus, the C. elegans primary small RNA pathway contains endogenously
produced ERGO-1–class 26G-RNAs, ALG-3/4–class 26G-RNAs (which bind the
redundant ALG-3 and ALG-4 paralogs), and PRG-1–associated 21U-RNAs, as well as
the exogenously sourced RDE-1–associated RNAs. C. elegans secondary small RNA
pathways include endogenously produced CSR-1–class 22G-RNAs and primary sRNA-
triggered WAGO–class 22G-RNAs (Ketting 2011). Despite some overlap, the biogenesis
and downstream targets are also defining features of each siRNA class, as discussed
below and outlined in Figure 1.
6

Figure 1. The C. elegans small interfering RNA pathway.
Graphical summary of different small RNA classes and interactions that comprise the siRNA
pathway
7
Primary sRNA Pathways
Primary endo-siRNAs: ERGO-1– and ALG-3/4–class 26G-RNAs
ERGO-1– and ALG-3/4–class siRNA production requires a multiprotein complex
called the ERI-complex, named for an Enhanced RNAi phenotype in mutants. The ERI-
complex associates with mRNA transcripts and uses the RNA-dependent RNA
polymerase (RdRP), RRF-3, to produce a complementary dsRNA (Simmer et al. 2002;
Lee et al. 2006; Gent et al. 2009). The mechanism by which the ERI-complex initially
recognizes a target transcript for silencing is still unclear, but the ERGO-1 and ALG-3
Argonautes are each required for production and stability of their respective siRNAs
(Conine et al. 2010). The resulting RRF-3–synthesized dsRNA undergoes further
processing by the lone C. elegans homolog of Dicer, DCR-1, an RNase-III ribonuclease
which operates in both siRNA and miRNA pathways (Ketting et al. 2001; Bernstein et al.
2001). DCR-1 cleaves the dsRNA into the class-defining 26-nucleotide long fragments
beginning with a 5’ guanine (G) (Ruby et al. 2006). Also as a result of DCR-1 cleavage,
26G-RNAs have a 5’ monophosphate and overhanging nucleotides which act as an
anchoring site for subsequent ERGO-1 or ALG-3/4 Argonaute binding (Pak and Fire
2007; Ma et al. 2004; Fischer et al. 2011). ERGO-1–associated siRNAs target deleterious
transcripts such as transposons and gene duplications in oocytes, embryos, and somatic
cells and ALG-3/4–associated siRNAs are enriched in spermatogenic cells and essential
for male fertility at elevated temperatures (Conine et al. 2010; Fischer et al. 2011).

Primary piRNAs: PRG-1–associated 21U-RNAs
In contrast to ERGO-1– and ALG-3/4–class siRNA production, the processing of
piRNAs is independent of DCR-1 activity. piRNAs, also known as 21U-RNAs, are instead
8
expressed as discrete transcriptional units from piRNA clusters: large, defined loci on
chromosome IV that contain individual piRNA sequences and promoters (Ruby et al.
2006; Das et al. 2008). The single-stranded, mature piRNAs contain a 5’ monophosphate
uridine, a 3’-2’-OMe, and are typically 21 nucleotides long, indicating posttranscriptional
processing, trimming, and protection from degradation (Ruby et al. 2006; Montgomery et
al. 2012). A recently described multi-protein complex named PETISCO is implicated in
piRNA 5’ end processing, and the exonuclease PARN-1 further trims the RNA to the
proper length in coordination with HENN-1 for 3’ methylation (Montgomery et al. 2012;
Tang et al. 2016; Rodrigues et al. 2019). These 21U-RNAs are predominantly expressed
in both hermaphrodite and male germlines, where they depend on and associate with the
PIWI–clade Argonaute PRG-1 to target many endogenous genes, including the abundant
Tc3 transposons (Batista et al. 2008; Das et al. 2008; Lee et al. 2012; Svendsen and
Montgomery 2018).

Primary exo-siRNAs: RDE-1–associated RNAs
Exogenous RNAi pathways can be initiated via laboratory manipulation and
introduction of dsRNAs, or via natural sources of dsRNA, such as viral infection. In the
laboratory, dsRNAs can be directly injected into the body cavity, gut, or germline of the
nematode, or C. elegans can be soaked in dsRNA for 24 hours, or, the most convenient
and widely used method, dsRNA can be fed to C. elegans in the form of dsRNA-
expressing bacteria (Fire et al. 1998; Tabara 1998; Timmons and Fire 1998; Timmons et
al. 2001; Kamath et al. 2000; Kamath et al. 2003a). For each method of artificial
exogenous RNAi initiation, long dsRNA spreads throughout the organism via the
9
transmembrane channel protein SID-1, for systemic RNA interference-deficient, and is
cleaved by DCR-1 in complex with the RNA-binding protein RDE-4, named for the RNAi
defective phenotype of mutants (Knight and Bass 2001; Tabara et al. 2002; Winston
2002). Exo-siRNAs sourced from viral infection require DCR-1, RDE-4, and an additional
DEAD-box RNA helicase, Dicer-related helicase DRH-1 for processing (Lu et al. 2009;
Ashe et al. 2013). Exogenous dsRNA as short as 38 base pairs can initiate RNAi, but
longer dsRNA is more efficiently processed by DCR-1 into 21-23 nucleotide long primary
exo-siRNAs with no 5’ nucleotide bias (Elbashir et al. 2001; Ashe et al. 2013). The
resultant exo-siRNAs, whether from the lab or from viral sources, are subsequently
loaded onto the Argonaute RDE-1 to initiate target silencing (Tabara et al. 1999; Yigit et
al. 2006). The biological function of exo-siRNAs behaves as a defense to viral infection,
but artificially induced exo-siRNAs can target any sequence as defined by the injected,
soaked, or fed dsRNA, a trait which has been widely co-opted for genetic manipulation
and study (further discussed on page 17).
A sRNA loaded onto its cognate Argonaute creates the RNA-induced silencing
complex (RISC). It is still unclear how the different small RNA classes are properly routed
to their Argonaute partners, but the downstream effect of RISC targeting to transcripts is
dictated by the Argonaute component. Most primary RISC Argonautes initiate
downstream production of an abundant class of secondary siRNAs.

Secondary siRNA Pathways
As first alluded to by Fire et al. (1998), amplification of small RNAs is required to
produce a widespread and heritable silencing signal. Downstream small RNA
amplification is achieved by the activity of two partially redundant RdRPs, EGO-1 and
10
RRF-1, which synthesize a distinct, highly abundant class of single-stranded 22-
nucleotide, 5’ guanine, and 5’ triphosphate-modified siRNAs, termed 22G-RNAs
(Smardon et al. 2000; Sijen et al. 2001; Aoki et al. 2007; Gu et al. 2009). Similar to primary
sRNA nomenclature, 22G-RNAs are further defined by the Argonaute with which they
associate. The major 22G-RNA classes include a general WAGO-class, which associate
with Worm-specific Argonautes, and CSR-1–class, which bind the CSR-1 Argonaute,
named for the chromosome-segregation and RNAi-deficient mutant phenotype (Yigit et
al. 2006; Claycomb et al. 2009). Adding to an already complex system of primary and
secondary siRNAs, CSR-1–class 22G-RNAs are produced endogenously, whereas
WAGO–class 22G-RNA amplification can be triggered by ERGO–class 26G-RNAs, exo-
dsRNA, some piRNAs, or even by other 22G-RNAs (Ketting and Cochella 2020). Though
some proteins components are shared, the molecular requirements for amplification vary
by pathway.

Secondary endogenous CSR-1–class 22G-RNAs
CSR-1–class siRNAs are a unique, poorly understood, and perhaps noncanonical
case of secondary siRNAs. CSR-1–class 22G-RNAs are produced endogenously via
EGO-1 activity. Though their 22-nucleotide structure and 5’ guanine bias is similar to other
secondary siRNAs, the mechanisms behind upstream initiation or possible de novo
targeting of EGO-1 to transcripts for CSR-1–class siRNA synthesis are still unclear
(Claycomb et al. 2009). CSR-1 targets include a wide array of germline genes and
possibly a subset of ALG-3/4 targets (Conine et al. 2013). Paradoxically, CSR-1 targets
appear to be protected instead of silenced: in csr-1 mutants, most target transcripts are
11
downregulated instead of displaying aberrant upregulation (Wedeles et al. 2013). This
gene licensing mechanism is hypothesized to counteract the widely-targeting 21U-RNAs
(Seth et al. 2013; Shen et al. 2018), but how these pathways interact to monitor proper
gene expression in the germline remains unknown. Regardless of a murky biogenesis
and mysterious mechanism, CSR-1–class siRNAs are required for fertility (Yigit et al.
2006).

Secondary WAGO–class 22G-RNAs
WAGO–class siRNAs depend on the RdRP RRF-1 for somatic and germline
amplification, but in the case of rrf-1 mutations, EGO-1 appears to compensate in the
germline (Gu et al. 2009). Primary Argonautes ERGO-1, PRG-1, and RDE-1 direct which
target transcripts are templated for RdRP production of secondary siRNAs, with
assistance from the Dicer-related helicase, DRH-3 (Duchaine et al. 2006; Nakamura et
al. 2007; Aoki et al. 2007). The resulting 22G-RNAs are antisense to the templated
transcript and exhibit spreading to regions both upstream and downstream of the original
RISC pairing site (Sijen et al. 2001; Pak and Fire 2007; Lee et al. 2007).
In addition to RdRPs, production of WAGO-class 22G-RNAs explicitly requires a
suite of proteins from the mutator (mut) family, which interact together to form the mutator
complex. Mut genes were originally discovered in an EMS mutagenesis screen which
isolated mutations that activated the Tc1 transposon (Collins et al. 1987). Subsequent
mutagenesis and RNAi screens identified the mut genes at the molecular level (Tabara
et al. 1999; Vastenhouw et al. 2003), but much remains to be understood about mutator
protein function and assembly. The nucleotidyl transferase MUT-2/RDE-3 is a key
12
mutator protein recently shown to add long 3’ poly-UG (pUG) tails to transcripts targeted
by RISC (Shukla et al. 2020). Addition of pUG tails also requires the endonuclease RDE-
8, which may cleave the RNA target to allow MUT-2 nucleotidyl activity (Tsai et al. 2015;
Shukla et al. 2020). pUG-ylation is hypothesized to mark RNA for secondary siRNA
synthesis, as it also recruits RRF-1 to the template transcript (Shukla et al. 2020). The
other mutator proteins are less understood: MUT-7 is a 3’-5’ exoribonuclease with
homology to human RNaseD and interacts with RDE-2/MUT-8 which contains no known
functional motif (Ketting et al. 1999; Tops et al. 2005). MUT-15/RDE-5 also contains no
known functional motifs (Tops et al. 2005). MUT-14 is a conserved DEAD-box helicase
with overlapping functions to its paralog, SMUT-1, both of which are required for germline-
specific RNAi (Tijsterman et al. 2002; Phillips et al. 2014). And finally, MUT-16 is a
nematode specific Q/N-rich protein which was anticipated to mediate protein-protein
interactions (Zhang et al. 2011). The exact function of MUT-16 within the mutator complex
was unknown until the work of Carolyn Phillips (further discussed on page 32) (Phillips et
al. 2012). Mutations in any individual mut genes disrupt RNA interference and cause
severe loss of WAGO-class 22G-RNAs (Gu et al. 2009; Gent et al. 2010; Vasale et al.
2010; Zhang et al. 2011; Phillips et al. 2012; Phillips et al. 2014; Tsai et al. 2015). Mutator
mutants also exhibit a reduction in ERGO-1–class 26G-RNAs due to loss of 22G-RNAs
targeting the sensor of siRNA, sosi-1, and eri-6[e-f]. siRNA repression of eri-6[e-f] is
necessary to promote production of ERI-6/7, an ERI-complex component specifically
required for synthesis of ERGO-1–class primary siRNAs (Zhang et al. 2011; Rogers and
Phillips 2020a). Resulting from the loss of key siRNAs, mutator mutants are temperature-
sensitive sterile, display increased frequencies of chromosome nondisjunction, have de-
13
repressed transposons, and are RNAi defective (Collins et al. 1987; Tabara et al. 1999;
Ketting et al. 1999).
siRNA amplification provides a solution to limited quantities of target transcript. For
endogenous pathways, amplification and maintenance of 22G-RNAs ensures siRNAs
can facilitate downstream silencing throughout the germline and are abundant enough to
be deposited in developing oocytes. For exogenous RNAi, amplification is crucial for a
robust silencing signal to be spread throughout the nematode and passed down to
progeny as a “memory” of encountered challenges (Fire et al. 1998; Grishok et al. 2000;
Lev et al. 2019).

RISC Assembly
As previously described, both primary and secondary siRNAs are loaded onto
Argonaute proteins, creating the actual effectors of RNAi and downstream function, RISC.
Though this nomenclature is more popular for other model organisms, here it is used as
a general term to refer to any Argonaute-sRNA unit. Among the 26 putative C. elegans
Argonautes, 19 have been confirmed to be expressed under normal physiological
conditions (Claycomb J.M., personal communication), yet there is incomplete
understanding of how multiple classes of sRNA are properly matched and loaded onto
their respective Argonaute. One hypothesis is that the sRNA sequence and features, such
as 5’ nucleotide bias, full or partial sequence complementarity, and 3’ modification,
generate favorable binding with some Argonautes but not others (Jannot et al. 2008;
Czech and Hannon 2011). The conserved binding pocket in Argonautes is created by the
PAZ domain, which binds the 3’ terminus with preference for nucleotide overhang and
modification, and the MID domain, which docks the 5’ terminus with specificity for 5’
14
nucleotide bias (Ma et al. 2004; Frank et al. 2010; Boland et al. 2010; Montgomery et al.
2012). The PAZ-MID binding pocket positions the siRNA along a conserved PIWI domain,
which contains an RNase-H like fold with a catalytically active DDH-motif in primary
Argonautes (Yigit et al. 2006). Lastly, a variable N-terminal region assists in unwinding
small RNA duplexes during loading (Kwak and Tomari 2012). Another idea for accurate
RISC assembly is that sRNA biogenesis binding factors, such as RDE-4 in the exo-RNAi
pathway, aid in recognition or guidance of proper Argonaute binding (Tabara et al. 2002;
Jannot et al. 2008). A final emerging mechanism posited for RISC assembly and sRNA
routing is that spatiotemporal Argonaute expression and sub-cellular localization
promotes a physical interaction to drive assembly or creates a barrier to prevent aberrant
RISC assembly (Ketting and Cochella 2020). These multiple mechanisms likely work in
combination to drive proper RISC assembly (Hutvagner and Simard 2008; Meister 2013).
At Argonaute binding, 26G-RNAs and exo-siRNAs are double stranded and must
become single stranded to allow for subsequent hybridization to a complementary target
transcript. In part, strand selection is dictated by strand asymmetry which predisposes
one strand, termed “passenger strand”, to be released or cleaved, and the other “guide”
strand to remain bound to the Argonaute. The guide strand results from the more
thermodynamically unstable 5’ sequence, which allows it to be more readily unwound and
loaded onto the Argonaute (Khvorova et al. 2003; Kwak and Tomari 2012). For some
Argonautes, including RDE-1, removal of the passenger strand can occur by direct
cleavage in the catalytically active RNase-H fold of the PIWI domain (Matranga et al.
2005; Steiner et al. 2008; Fischer et al. 2011). The outcome of RISC assembly is an
15
elegant modular system in which the siRNA sequence specifies a complementary target
and the bound Argonaute dictates a downstream action.

Mechanisms of Action
All primary Argonautes contain the catalytically active DDH motif in the PIWI
domain RNase-H fold, yet do not directly cleave the target transcripts (Yigit et al. 2006).
Instead, primary Argonautes facilitate cleavage through endonuclease recruitment. This
has been explicitly demonstrated for exo-siRNAs in which RDE-1 recruits RDE-8 to
cleave the target, thereby initiating pUGylation by MUT-2/RDE-3 and marking it for
mutator-dependent 22G-RNA synthesis (Tsai et al. 2015; Shuckla et al. 2020). Primary
Argonautes may also remain bound to the target to inform downstream function.
Therefore, the main outcome of primary Argonautes in the siRNA pathway is to initiate
downstream 22G-RNA synthesis.
Secondary WAGO–class Argonautes have lost the enzymatic DDH motif and are
therefore unable to directly cleave target RNA. So how do secondary Argonautes enact
RNA silencing? The mechanisms of downregulation can be broadly categorized as either
transcriptional or post-transcriptional. Transcriptional silencing occurs in the nucleus and
is carried out by the nuclear WAGO–class Argonautes HRDE-1 (heritable RNAi deficient)
and NRDE-3 (nuclear RNAi defective), which direct the deposition of H3K9Me3
repressive chromatin marks on the genomic locus of the target transcript in the germline
and soma, respectively (Burton et al. 2011; Gu et al. 2012; Buckley et al. 2012; Ashe et
al. 2012; Spracklin et al. 2017). Germline transcriptional silencing also requires HRDE-2,
which facilitates binding of siRNA cofactors for HRDE-1 targeting and function, and
WAGO-4, which transfers 22G-RNAs from parents to progeny to perpetuate the silencing
16
signal in the following generation (Spracklin et al. 2017; Xu et al. 2018). The resulting
transcriptional silencing is able to be inherited across multiple generations, providing the
nematode with a “memory” of acquired silencing signals, termed RNAi inheritance
(Grishok et al. 2000; Vastenhouw et al. 2006; Alacazar 2008; Lev et al. 2019; Houri-Zeevi
et al. 2020). WAGO silencing via cytoplasmic post-transcriptional mechanisms is less
understood, but is presumed to occur by the recruitment of exonucleases or the inhibition
of translational complexes ultimately destabilizing the transcript in a similar mechanism
of action as miRNA Argonautes (Hutvagner and Simard 2008). The resulting gene
silencing and repression are highly efficient.

RNAi Technologies and Tools
Exogenous RNAi can silence a target to biochemically undetectable levels,
effectively creating a genetic knockdown. This feature, paired with the ease of
administering dsRNA via bacterial feeding, renders RNAi a highly useful molecular
technology which can be used for reverse genetics studies in C. elegans and other
organisms. Two large and commercially available libraries contain RNAi clones targeting
the majority of annotated C. elegans genes, allowing for widespread and high-throughput
genetic analysis (Kamath and Ahringer 2003b, Rual et al. 2004). Bacterial RNAi clones
can also be made to any desired RNA by inserting the sequence of interest to a plasmid
containing two antiparallel T7 promoters, resulting in dsRNA expression (Timmons et al.
2001). Results are fast: RNAi silencing can be phenotypically observed as early as 24
hours-post feeding (Kamath et al. 2000). In addition to its utility as a molecular tool, RNAi
is being explored as a clinical therapeutic. Target specificity, utilization of endogenous
cellular machinery, and potent downregulation make RNAi an attractive avenue for acute
17
disease treatment and clinical gene regulation, but there are some challenges in
introducing siRNA to patients’ cells which warrants further development of protective
siRNA delivery mechanisms (Kim et al. 2019). Despite the challenges, there have been
over 30 clinical trials of siRNA-based therapeutics, 40% of which are directed towards
cancer or cancer-related diseases (Kim et al. 2019). As of today, only three siRNA
therapeutics are FDA approved, but more are in phase 3 clinical trials with hopes of being
approved for treatment of a wide range of diseases (Zhang et al. 2021a). The evolutionary
conservation of general RNAi mechanisms from nematodes to humans, coupled with the
powerful and accessible RNAi response of C. elegans, places worms as an ideal model
organism with which to explore siRNA pathways.

Consequences of disruption
In C. elegans, the siRNA pathways are critical for health. Disruptions in shared
upstream elements of the pathways, such as dcr-1, results in developmental somatic
defects, sterile adults, and a failure to respond to exo-RNAi (Ketting et al. 2001). In the
piRNA pathway, mutations in the 21U-RNA processing protein, henn-1, cause sterility at
elevated temperatures and prg-1 mutant animals lose 21U-RNAs, fail to silence the Tc3
transposon, and are also temperature-sensitive sterile (Batista et al. 2008; Das et al.
2008; Montgomery et al. 2012). Mutations in rde-4 of the exo-RNAi pathway also result
in temperature-sensitive sterility, developmental defects, and a failure to engage RNAi or
respond to viral threats (Tabara et al. 1999; Tabara et al. 2002; Lu et al. 2009; Blanchard
et al. 2011). Curiously, animals with mutations in upstream components of the
endogenous 26G-RNA pathway, including the ERI-complex RdRP, rrf-3, mount an
enhanced response to exogenous siRNAs, such that fed dsRNA provokes a greater
18
accumulation of 22G-RNAs, sensitizing the nematode to neuronal RNAi and RNAi with
previously incomplete phenotypes (Kennedy et al. 2004). The enhanced RNAi phenotype
arises from reduced competition for molecular resources shared between the exo-RNAi
pathway and the 26G-RNA pathway (Lee et al. 2006). In addition to an enhanced RNAi
phenotype, ERI-complex mutants are temperature-sensitive sterile and have a reduced
brood size which results from loss of endogenous ERGO-class 26G-RNAs and
associated germline surveillance (Duchaine et al. 2006; Fischer et al. 2008; Fischer et al.
2011). Loss of alg-3/4 and the corresponding class of 26G-RNAs also causes reduced
brood size at permissive temperatures and complete sterility at elevated temperatures.
Because ALG-3/4–class 26G-RNAs mediate proper gene expression during male
gametogenesis, sterility is male-specific and manifests as arrested spermatocytes, lower
sperm counts, and defective spermatid activation (Conine et al. 2010).
The most widespread phenotype of primary sRNA and Argonaute loss is sterility
at elevated temperatures. Interestingly, the only Argonaute to cause acute sterility and
embryonic lethality at non-elevated temperatures is csr-1 (Yigit et al. 2006). As CSR-1–
class 22G-RNAs are thought to protect essential transcripts and are antisense to
thousands of germline-expressed genes, disruption of csr-1 causes widespread negative
phenotypes. csr-1 mutants display under-proliferated germlines, chromosome
segregation defects, and abnormal germ cell nuclei (Yigit et al. 2006; Claycomb et al.
2009).
However, the majority of primary RNA pathways funnel through mutator complex-
dependent amplification, and thus the loss of any mutator complex component results in
the loss of 22G-RNAs targeting over 2,000 genes and compounds the effects of ergo-1,
19
prg-1, rde-1 and other upstream pathway mutants (Zhang et al. 2011; Phillips et al. 2014).
Mutator mutants display sterility within one generation at elevated temperatures, but
display a low-penetrance larval arrest, affecting about 20% of offspring, and lay half as
many eggs as wild-type animals even at permissive temperatures (Collins et al. 1987;
Rogers and Phillips 2020b). Mutator mutants also show activated expression of
transposons, an inability to respond to exogenous RNAi, and chromosome nondisjunction
resulting in elevated male progeny (Collins et al. 1987; Ketting et al. 1999; Gu et al. 2009;
Zhang et al. 2011). Together, primary and secondary siRNAs provide protection against
selfish genetic elements and viral threats, and ensure proper gene expression during
gametogenesis and embryogenesis, making small RNA pathways a crucial element of C.
elegans health.

SMALL RNA PATHWAY ORGANIZATION
With so many health and developmental outcomes relying on intertwined and
complex siRNA pathways, an outstanding question remains: How are the siRNA
pathways organized to facilitate proper gene regulation and ensure robust silencing? This
inquiry encapsulates numerous facets of the siRNA pathway, including: (1) how are the
different classes of siRNAs routed to proper Argonaute partners?, (2) how are nascent
transcripts monitored so sRNA biogenesis machinery can identify deleterious RNAs?, and
critically (3) how, physically, do the components of the small RNA pathways recognize
and interact with one another in a crowded cytoplasmic space to ensure efficient
silencing? These remaining questions are only beginning to be answered and create the
niche in which my doctoral studies push the current boundary of knowledge.
20

P granules: Early Clues to siRNA Organization
P Granule Discovery
The first clue to shed light on the organization of RNAi pathways was the discovery
that many small RNA components are subcellularly localized to P granules, the C.
elegans germ granule. Germ granules are a highly conserved feature of development —
found in over 80 species across the animal kingdom — that describes membraneless,
cytoplasmic organelles which localize to, or are found in, germ cells (Eddy 1975; Schisa
2012). Evidence of the membrane-less nature of germ granules is even prominent in the
nomenclature of early literature, where the structures are described as nuage, from the
French meaning “cloud” (Andre and Rouiller 1957). In the broadest sense, germ granules
function to inform cell fate, maintain germ cell totipotency, or protect fertility, and are
ultimately required for germ cell function (Voronina et al. 2011).
The C. elegans germ granule, P granules, were first fluorescently visualized by
Susan Strome in a serendipitous late-night “eureka” moment. A rabbit-anti mouse IgG
secondary antibody cross-reacted with P granules, revealing fluorescent globules that
partitioned exclusively to the embryonic P-lineage germline precursor cells (Strome and
Wood 1982). This discovery evoked such a great excitement in Strome that she
immediately shared her novel results with the closest person; thus, the second observer
of fluorescent P granules was the University of Colorado Boulder’s night-shift janitor
(Maartens 2018).

21
P Granule Localization and Composition
The discovery of P granule antibodies proved a great asset, and researchers
began to probe the localization, composition, and function of the C. elegans germ granule.
P granules are maternally deposited into the zygote, where they initially localize as puncta
throughout the cytoplasm. Just prior to the first asymmetric cleavage, P granules partition
to the posterior pole and are asymmetrically sequestered in the P1 blastomere upon
cleavage. P granules continue to asymmetrically partition with the subsequent P-lineage
daughter blastomeres P2, P3, and P4, where they increasingly associate with the nuclear
periphery and grow in size, occasionally reaching as large as 4 µm in diameter (Strome
and Wood 1982, Brangwynne et al. 2009; Updike and Strome 2010). In embryos, the P4
cell divides once more to produce the Z2 and Z3 progenitor germ cells, which only
continue development after hatching and feeding to eventually give rise to the adult
germline (Sulston et al. 1983). During gonadal development, P granules associate with
the nuclear membrane of all larval and adult germ cells, except nuclei undergoing
apoptosis or spermatogenesis (Sheth et al. 2010).
About 75% of nuclear pores interact with P granules, meaning the majority of
nascent transcripts pass through P granules before export to the cytoplasm (Pitt et al.
2000; Sheth et al. 2010). Both nuclear pore proteins and P granule components are
required for this interaction, which is mediated through the FG-repeat domains in nuclear
pore complex proteins and the FG-repeat-rich N-terminal region of GLH-1, a conserved
Vasa-related DEAD-box helicase that is a constitutive P granule protein (Roussell and
Bennett 1993; Schisa et al. 2001; Updike and Strome 2009; Updike et al. 2011). GLH-1
is one of 4 Vasa-related helicases (GLH-1–4) that localize to P granules and both GLH-1
22
and GLH-2 are individually required for fertility (Gruidl et al. 1996). Other essential
constitutive components of P granules are the RGG-domain containing PGL-family
proteins, PGL-1 and PGL-3. PGL-1 is required for proper germline development and
fertility at elevated temperatures and pgl-1; pgl-3 double mutants display fertility defects
in all temperature ranges (Kawasaki et al. 1998; Kawasaki et al. 2004). PGL-1 depends
on GLH-1 for assembly at the nuclear periphery (Kawasaki et al. 1998), and glh-1 mRNA
and protein accumulation depend on a nematode-specific protein, DEPS-1, for defective
P granules and sterile (Spike et al. 2008; Updike and Strome 2010). As the name
suggests, P granules are lost in deps-1 mutants (Spike et al. 2008). Loss of P granules,
or the combinatorial depletion of key P granule components, causes errant accumulation
of somatic transcripts in the hermaphrodite germline and eventual reprogramming
towards soma: neurons and muscle cells (Updike et al. 2014; Knutson et al. 2017). From
the observations that P granules interact with nascent transcripts, and loss of P granules
causes differentiation of germ cells into somatic cells, P granules are hypothesized to be
RNA processing centers which monitor gene expression to ensure germ cell identity.

siRNA Pathway Proteins Localize to P granules
The model of P granules as RNA processing centers is bolstered by discoveries
that key siRNA pathway components localize to germ granules — a finding which is also
true for Drosophila, zebrafish, and mice germ granules (Lim and Kai 2007; Brennecke et
al. 2007; Houwing et al. 2007; Aravin et al. 2009). In C. elegans, proteins from all aspects
of the RNAi pathway are localized to P granules, including small RNA processing and
biogenesis factors such as PARN-1, DRH-3, and EGO-1, as well as Argonautes PRG-1,
23
ALG-3, CSR-1, and WAGO-1 (Batista et al. 2008; Gu et al. 2009; Claycomb et al. 2009;
Conine et al. 2010; Tang et al. 2016). These discoveries opened two new tantalizing and
intertwined hypotheses: P granules are not just RNA processing centers, but RNA
surveillance centers, and small RNA pathway components are organized, at least in part,
by a membraneless organelle.

P Granules Have Liquid-Like Properties
Around the same time that P granule and small RNA protein colocalization was
being investigated, the canonical ideas of subcellular organization primarily revolved
around membrane-bound organelles, which are divided from the cytosol by a
phospholipid bilayer and provide discrete environments suited for particular processes.
For example, the endoplasmic reticulum provides an oxidizing environment to facilitate
protein folding, whereas the lysosome maintains an acidic environment for protein
destruction (Margittai et al. 2015; Luzio et al. 2007). However, P granules, like other germ
granules, appeared to be membraneless. At the molecular level, how does the
membraneless P granule create a distinct compartment to sequester proteins, capture
RNA, and act as an mRNA surveillance center?
The answer to this enigma came in 2009; when P granules were shown to have
liquid-like properties. Cliff Brangwynne and colleagues published the findings that P
granules (1) asymmetrically localize to the P-cell lineage during early embryogenesis via
protein dissolution and condensation, (2) drip off nuclei when a force is applied and
subsequently fuse together to create a larger, round droplet, and (3) exhibit rapid internal
molecular rearrangements as determined through fluorescence recovery after
24
photobleaching (Brangwynne et al. 2009). Simply put, P granules behave similarly to oil
droplets in water. In biological terms, a concentrated protein phase surrounded by a bulk
cytoplasmic phase creates the defined, membraneless boundary of P granules through a
process called phase separation.

Phase Separation
Early Ideas of Phase Separation
Perhaps the earliest concepts of biological phase separation originated with Carl
Nägeli’s study of the organization and structure of starch granules in plant cells, work that
would later be referred to as micellar theory (Scott 1891). Later, in eukaryotic cells, phase
separation revealed itself as “suspended drops… of different chemical nature” and
“protoplasm… composed of innumerable minute granules, or microsomes, suspended in
a clearer, less deeply staining, continuous substance” as proposed by Edmund Wilson in
1899 when observing the cytoplasm of sea star and urchin eggs (Wilson 1899).
Phase separation continued to be studied in the fields of chemistry and polymer
physics, but the idea of biomolecular phase separation lost popularity after the discovery
of protein structures — alpha helixes, beta sheets — in the x-ray crystallography era of
protein study (Eisenberg 2003; Hyman and Simons 2012). The discovery of the liquid-like
nature of P granules launched the field of biomolecular phase separation back into
popularity and spurred the investigation, characterization, and experimental validation of
many additional phase-separated droplets. P granules and other structures with similar
liquid-like properties were referred to as droplets, membraneless organelles, or
condensates (Shin and Brangwynne 2017). Biological phase separation quickly became
a new paradigm with which to view cellular organization.
25

Properties of Phase Separation
A few key properties define biomolecular phase-separated condensates. First,
phase-separated condensates are concentration dependent, assembling only above a
certain threshold concentration, Csaturation, wherein molecules are abundant enough to
interact and coalesce (Li et al. 2012; Weber and Brangwynne 2015). While many proteins
can be forced to undergo phase separation in in vitro laboratory settings (as is necessary
for protein crystallography), membraneless organelles condense at physiological or near-
physiological conditions (Alberti et al. 2019). This threshold of condensation can be
altered via salt concentration, RNA concentration, pH, post-translational modification, and
temperature, making coacervation and dissolution susceptible to the surrounding
environment (Brangwynne et al. 2009; Wang et al. 2014; Molliex et al. 2015; Elbaum-
Garfinkle et al. 2015; Kroschwald et al. 2018; Dignon et al. 2020). This property is often
described using a phase diagram in which protein concentration is plotted against the
altering factor (temperature, salt concentration, etc) to create a binodal curve, under
which two phases exist (Alberti et al. 2019). The ability to respond to changing
environments constitutes a second characteristic of condensates, and provides a means
by which cellular environments can promote or prevent protein condensation. Lastly, the
liquid-like nature of membraneless organelles is revealed in the ability of molecules to
rearrange rapidly within the condensate and exchange readily with the bulk phase (Phair
and Misteli 2000; Brangwynne et al. 2009; Sheth et al. 2010; Molliex et al. 2015). This
property is often probed with fluorescent recovery after photobleaching (FRAP), which
monitors the rate of diffusion of fluorescently-tagged proteins in repopulating a
26
photobleached region. Phase separated proteins can quickly repopulate a photobleached
region on a scale ranging from a few seconds to a few minutes (Brangwynne et al. 2009;
Feric et al. 2016).

Composition of Membraneless Organelles
Condensates gain these biophysical properties from the proteins and biomolecules
with which they form and interact. Phase separated condensates are often
heterogeneous, containing multiple interacting protein species and/or RNAs. Proteins that
phase separate often contain low complexity sequences or intrinsically disordered
regions, characterized by structural flexibility and the ability to sample multiple
conformations (Lin et al. 2015; Babu 2016). Functionally, intrinsically disordered regions
are promiscuous binding partners and can serve as a scaffold to assemble multi-protein
complexes (Haynes et al. 2006; Protter et al. 2018). Intrinsically disordered regions
typically contain charged or polar amino acids that facilitate multiple weak, electrostatic
interactions, which are key drivers of phase assembly (Pak et al. 2016; Dignon et al.
2020).
Many different chemical interactions contribute to phase assembly and inform the
unique conditions under which proteins condense or disassemble (Dignon et al. 2020). In
addition to charge-charge interactions of polar amino acids, aromatic residues such as
Tyrosine contribute to phase separation via π-π or cation-π interactions (Lin et al. 2017;
Li et al. 2018; Vernon et al. 2018). These short, interacting motifs within disordered
proteins have been termed “stickers”, which are interspersed by longer, flexible, non-
interacting regions called “spacers” in a stickers-and-spacers model of multivalent
27
interactions (Choi et al. 2020). In another interpretation of the stickers-and-spacers
model, a protein can have structured “sticker” regions, which facilitate protein- or RNA-
binding, and disordered “spacer” regions, which provide flexible linkers and phase-
stabilizing interactions. The patterning and frequency of stickers versus spacers provides
a distinctive condensation profile to each protein (Choi et al. 2020). Overall, condensate
formation is two-fold: (1) protein-protein or protein-RNA molecules polymerize via
specific, stronger multivalent interactions, such as protein or RNA binding, and (2) these
polymers de-mix from the bulk phase into a condensate, which is stabilized by many
weak, multivalent, and hydrophobic interactions (Lin et al. 2017).
Assembly and disassembly of condensates can occur rapidly through destabilizing
multivalent interactions, such as blocking π-π aromatic interactions via phosphorylation
of Tyrosine, or reducing cation-π interaction through dimethylation of Arginine (Wang et
al. 2014; Lin et al. 2017; Ryan et al. 2018). For example, asymmetric P granule
disassembly is controlled by phosphorylation in early embryos (Wang et al. 2014).
Additionally, 1,6-hexanediol is an aliphatic alcohol commonly used in experiments to
destabilize weak, hydrophobic interactions and dissolve condensates (Kroschwald et al.
2017). The readily tunable nature of condensates positions them as versatile functional
organelles within cells.

Functionality in Cells
The functional capacities of condensates are expansive, but can be broadly
categorized into three actions. First, and perhaps most intuitively, condensates can
sequester or release proteins. This action can create biochemical reaction crucibles,
28
provide sensing or buffering capacities, or enable signal transduction. Second, multivalent
interactions within droplets can form a nanoscale mesh network which acts as a complex
biomolecular filter (Updike et al. 2011; Nott et al. 2016). Lastly, and perhaps as a
combination of the two prior functionalities, different phases can provide the framework
within which cellular pathways are organized (Shin and Brangwynne 2017; Alberti et al.
2019). One striking example of functional organization within phases is the three-tiered,
interior-to-exterior arrangement of ribosome biogenesis and assembly factors within the
nucleolus, where ribosomal RNA is transcribed in the innermost phase, processed in the
intermediate phase, and assembled onto ribosomal proteins in the outermost phase
(Boisvert et al. 2007; Feric et al. 2016).
As the field grows, phase separation is being uncovered as an essential and near-
ubiquitous strategy of cellular function. Phase separation occurs in organisms ranging
from bacteria, where condensation coordinates RNA Polymerase clusters and gene
expression (Azaldegui et al. 2020), to tardigrades, where vitrification protects cells from
total desiccation and allows the animal to live for over a decade in a hardened, dormant
state (Boothby et al. 2017), to humans, where phase separation of heterochromatin
mediates gene silencing (Larson et al. 2017).

Consequences of Phase Separation
Despite widespread functions and near-ubiquitous nature of condensates, the
strategy of phase separation is not without consequence. While the process of forming
membraneless organelles is highly tunable via protein and RNA constituents, surrounding
environment, and post translational modification, this regulatory process also has
29
opportunity to go awry. Liquid-like condensates can progress to gelation and non-
reversible aggregation under certain conditions, including mutations and misexpression
(Shin and Brangwynne 2017). Irreversible aggregates are toxic to cells and can ultimately
cause cell death (Bolognesi et al. 2016). Studies of the human intrinsically disordered
RNA-binding protein, Fused-in-sarcoma (FUS), have shed light on underlying causes of
the progressive neurodegenerative disease, Amyloid Lateral Sclerosis (ALS); Patient-
derived mutations in FUS can cause pathological aggregation in cells, resulting in cell
death (Patel et al. 2015). New insights to the regulation and functions of phase separation
can benefit disease biology.

Phase Separation and Cellular Organization
Fundamentally, it is becoming more and more evident that biomolecular
condensates bridge the gap between ångström-scale molecules and micrometer-scale
protein assemblies visible by microscopy (Li et al. 2012). Phase separation can impart
organization to a cell crowded with different pathways, signaling molecules, and proteins,
but this intersection of phase separation and pathway organization offers much to be
explored.

siRNA Pathways are Organized Within Multiple Phase-separated Compartments
Z Granules
The connection between small RNA proteins and phase separated P granules
delivered a promising area of study for the organization of sRNA pathways and was soon
bolstered by the discovery of an additional phase-separated small RNA pathway
condensate: Z granules, named after the first identified component, a Zinc finger NFX1-
30
type homolog, ZNFX-1 (Wan et al. 2018; Ishidate et al. 2018). Z granules first appear
colocalized with P granules in early embryos, but de-mix around the 100-cell stage to
form separate and adjacent granules at the nuclear periphery (Wan et al. 2018).
Consistent with characteristics of phase separation, Z granules undergo rapid molecular
rearrangement and have a tunable viscosity, which is regulated by the piRNA-induced
silencing defective/Z granule surface protein PID-2/ZSP-1 (Wan et al. 2018; Wan et al.
2021; Placentino et al. 2021). Notably, Z granules also contain WAGO-4, a secondary
Argonaute required for transmission of secondary siRNAs to progeny (Wan et al. 2018;
Xu et al. 2018). Indeed, znfx-1 mutants respond normally to RNAi, but are unable to pass
the silencing signal on to progeny (Wan et al. 2018). ZNFX-1 also directly interacts with
a number of small RNA proteins, including EGO-1 and CSR-1, and interacts with WAGO-
1 and PRG-1 in a partially RNA-dependent manner (Ishidate et al. 2018). Finally, ZNFX-
1, in coordination with other factors, appears to balance the production of mutator-
dependent 22G-siRNAs across transcripts, preventing accumulation of siRNAs at the 5’
end of transcripts (Ishidate et al. 2018). Z granules, therefore, organize the
transgenerational inheritance of small RNA pathways and balance the production of
secondary siRNAs across transcripts.

Mutator Foci
Though key siRNA biogenesis and Argonaute proteins localize to liquid-like P
granules and Z granules, one mystery remained: how were the critical mutator complex
proteins organized and where did 22G-RNA amplification occur? Microscopy work by
Phillips et al. (2012) revealed that fluorescently-tagged mutator complex proteins localize
31
together and create punctate, perinuclear foci in the germline, termed Mutator foci. The
22G-siRNA-producing RdRP, RRF-1, also strongly localizes to Mutator foci (Phillips et al.
2012). Furthermore, some individual mutator components rely on a hierarchy of
requirements for recruitment to Mutator foci, such that MUT-7 requires RDE-2 for
localization (Tops et al. 2005), but the nucleation of all mutator components rely upon the
Q/N-rich protein MUT-16; mut-16 mutants are unable to form Mutator foci and lose the
vast majority of WAGO–class 22G-RNAs (Zhang et al. 2011; Phillips et al. 2012).
Curiously, Mutator foci localize directly adjacent to P granules, yet still appear structurally
distinct (Phillips et al. 2012). Even after dissolution of key P granule components by
simultaneous RNAi knockdown of glh-1 and glh-4, Mutator foci are still present,
suggesting a distinct structure of Mutator foci (Phillips et al. 2012). Finally, Z granules are
often observed to localize between P granules and Mutator foci (Wan et al. 2018).
Together, the dependency of 22G-RNA production on the mutator complex define Mutator
foci as siRNA amplification centers, and the adjacency of Mutator foci to P granules and
Z granules establish Mutator foci as an additional compartment in which siRNA pathways
are organized.

SIMR Foci
The most recently discovered compartment involved in sRNA pathways are SIMR
foci, of which only two protein localizations are currently known: the extended Tudor
domain protein SIMR-1 and the RNAi spreading defective, RSD-2 (Sakaguchi et al. 2014;
Manage et al. 2020). SIMR-1 was first identified in a MUT-16 immunoprecipitation, but
found to create functionally independent foci directly adjacent to Mutator foci. The name
32
SIMR-1 is derived from siRNA-defective and mortal germline, as simr-1 mutants lose
some mutator-dependent siRNAs and become sterile over multiple generations at
elevated temperature (Manage et al. 2020). Interestingly, simr-1 mutants desilence a
piRNA sensor and lose small RNAs specifically mapping to piRNA targets (Manage et al.
2020). This mutant phenotype, along with a conserved role of Tudor domain proteins in
both piRNA pathways and in protein-protein interactions (Pek and Kai 2012), leads to the
hypothesis that SIMR foci act in part to direct piRNAs or piRNA targets to Mutator foci for
downstream 22G-RNA synthesis (Manage et al. 2020). Furthermore, the colocalizing
factor RSD-2 acts in the exogenous siRNA pathway to ensure efficient RNAi, suggesting
that SIMR foci more generally mediate the transition between primary and secondary
small RNA pathways (Han et al. 2008; Sakaguchi et al. 2014; Manage et al. 2020).

Limits of Current Understanding
Thus, at least four distinct compartments — P granules, Z granules, SIMR foci,
and Mutator foci — constitute perinuclear nuage and are involved in RNA surveillance in
germ cells. Two of these compartments, Z granules and SIMR foci, were discovered only
within the past few years. The initial characterization of nuage compartments has
demonstrated individual contributions to siRNA pathway organization, but many
compartments lack extensive categorization of protein components and the specific
protein-protein interactions on which they rely for association. Additionally, the regulation
of compartment formation and requirements for nucleation have only been significantly
examined in P granules, but are lacking for Z granules, Mutator foci, and SIMR foci. Lastly,
the physical interaction and spatial orientation of all four compartments has only been
briefly explored with diffraction-limited microscopy. It is therefore unclear how biomolecule
33
exchange is facilitated between compartment boundaries, such that one protein is found
throughout multiple compartments. The current incomplete understanding of the C.
elegans nuage assemblage prevents formation of a comprehensive model to describe
how multiple compartments organize small RNA pathways and facilitate RNA silencing.

CREATING A SPATIAL MODEL OF RNA SILENCING
To begin to address how these multiple distinct compartments coordinate RNA
silencing, my doctoral work focuses on understanding Mutator foci within the context of
small RNA pathway organization. In my first chapter, I examine the protein-protein
interactions of MUT-16 and the biophysical properties of Mutator foci. I find that distinct
regions of MUT-16 nucleate interactions with other mutator complex proteins, adding
depth to understanding the localization requirements of mut-class proteins to the mutator
complex, and establish that the C-terminal region of MUT-16 is required for foci formation.
Importantly, I discover that Mutator foci have liquid-like properties and assemble via
phase separation (Uebel et al. 2018). Building upon the foundations of phase-separated
P granules and Z granules, I propose that siRNA organization is facilitated through
multiple phase-separated condensates.
To fully understand how Mutator foci behave as phase-separated condensates, I
probe the spatiotemporal regulation of MUT-16 and requirements for Mutator foci
formation. In my second chapter I demonstrate that Mutator foci first arise in 100-cell
embryos and persist throughout all larval stages. Furthermore, and in contrast to P
granules, I show that Mutator foci are also present in spermatids, implicating a role for the
34
mutator complex in paternal siRNA inheritance. Lastly, I determine that both RNA and the
meiotic cell cycle influence Mutator foci presence and morphology (Uebel et al. 2020).
Finally, I aim to contextualize Mutator foci as an organizing factor in coordination
with other phase-separated nuage compartments. In my last chapter, I use high resolution
microscopy to visualize the interaction between Mutator foci, P granules, Z granules, and
SIMR foci and discover discreet populations of nuage compartment assembly. I find that
P granule and Mutator foci interaction is independent of the germline environment when
ectopically expressed, dynamic, and able to be re-established after perturbation,
suggesting that cells may be able to coordinate the assembly and association of different
compartments to regulate RNA silencing. Finally, I discover a previously undescribed
toroidal P granule morphology, in which some P granules encompass Z granules, SIMR
foci, and Mutator foci with a consistent exterior-to-interior organization. I additionally find
that P granules associate with other nuage compartments in distinct populations,
suggesting a functional subdivision of different nuage assemblages.
Taken together, my doctoral work begins to answer the outstanding question of
how the components of siRNA pathways are physically organized within the cytoplasm to
accomplish efficient RNA silencing. I create a more accurate model of RNA silencing
through the lens of phase separation and reinforce that C. elegans nuage, comprised of
multiple phase-separated compartments, is a dynamic and critical organizer of RNA
surveillance.
35
CHAPTER 1: MUTATOR FOCI FORMATION
Mutator foci are phase-separated condensates nucleated by MUT-16

Associated Publication: Uebel C. J., D. C. Anderson, L. M. Mandarino, K. I. Manage, S.
Aynaszyan, et al., 2018 Distinct regions of the intrinsically disordered protein MUT-16
mediate assembly of a small RNA amplification complex and promote phase separation
of Mutator foci. PLoS Genet. 14: e1007542. https://doi.org/10.1371/journal.pgen.1007542

PREFACE
The groundwork for this project began before I joined the Phillips lab. Dr. Dorian
Anderson, a postdoc in the lab, launched this work by creating many of the strains and
capturing a few preliminary images. Because of this foundational strain creation, I was
able to begin conducting experiments immediately upon my joining in April 2017. This
project was attractive to me for two reasons. First, it would rely heavily upon microscopy,
a technique which has always fascinated me. The second reason was purely curiosity-
driven; I had never previously heard about biomolecular phase separation, as it was such
a novel concept that not even the newest undergraduate biology textbook covered it. At
the intellectual genesis of this project, Carolyn suspected that Mutator foci were phase-
separated condensates and this became a hypothesis I was determined to test.

ABSTRACT
In C. elegans, efficient RNA silencing requires small RNA amplification mediated
by RNA-dependent RNA polymerases (RdRPs). RRF-1, an RdRP, and other mutator
complex proteins localize to Mutator foci, which are perinuclear germline foci that
associate with nuclear pores and P granules to facilitate small RNA amplification. The
mutator complex protein MUT-16 is critical for Mutator foci assembly. By analyzing small
deletions of MUT-16, we identify specific regions of the protein that recruit other mutator
36
complex components and demonstrate that it acts as a scaffolding protein. We further
determine that the C-terminal region of MUT-16, a portion of which contains predicted
intrinsic disorder, is necessary and sufficient to promote Mutator foci formation. Finally,
we establish that MUT-16 foci have many properties consistent with a phase-separated
condensate and propose that Mutator foci form through liquid-liquid phase separation of
MUT-16. P granules, which contain additional RNA silencing proteins, have previously
been shown to have liquid-like properties. Thus, RNA silencing in C. elegans germ cells
may rely on multiple phase-separated compartments through which sorting, processing,
and silencing of mRNAs occurs.

INTRODUCTION
RNA silencing is an anciently conserved pathway that regulates gene expression in
most eukaryotes. Key to this pathway are members of the Argonaute protein family, which
bind a diverse set of small regulatory RNAs, ranging from ~18-30 nucleotides in length.
This small RNA-Argonaute complex regulates fully or partially complementary mRNAs at
the level of transcription, translation, and stability (Hutvagner and Simard 2008; Claycomb
2014). By regulating both endogenous and foreign RNAs, small RNAs maintain proper
gene expression, silence deleterious RNA products, and play critical roles in
development, chromosome segregation, transposon silencing, fertility, and viral defense
(Ketting 2011; Claycomb 2014).
Small RNAs can be generated through a variety of mechanisms. Long double-
stranded RNAs (dsRNAs), which can be produced from exogenous or endogenous
sources, are cleaved into small-interfering RNAs (siRNAs) by the RNase III-like enzyme
37
Dicer (Bernstein et al. 2001; Ketting et al. 2001). When these siRNAs are produced from
a primary dsRNA source, such as viruses or convergent transcription, they are referred
to as primary siRNAs. In many organisms, including C. elegans, plants, and fungi, RNA-
dependent RNA polymerases (RdRPs) use mRNAs targeted by primary siRNAs as a
template to amplify and maintain the silencing signal (Ghildiyal et al. 2009; Gu et al. 2009;
Vasale et al. 2010; Gent et al. 2010). RdRPs can function either by synthesis of additional
long dsRNA and coordinated cleavage by Dicer into secondary siRNAs, or, as in C.
elegans, RdRPs can directly synthesize secondary siRNAs antisense to the target mRNA
(Pak and Fire 2007; Lee et al. 2007; Colmenares et al. 2007).
In previous work, we characterized MUT-16 as a protein essential for RNA silencing
and nucleation of a nuclear pore-associated RNA silencing compartment in C. elegans
germ cells called Mutator foci (Phillips et al. 2012). MUT-16 recruits many other proteins
required for RNA interference and endogenous small RNA biogenesis to Mutator foci,
including the nucleotidyl transferase MUT-2, the 3’-5’ exonuclease MUT-7, the DEAD-box
RNA helicases MUT-14 and SMUT-1, and two proteins of unknown function, RDE-2 and
MUT-15 (Phillips et al. 2012; Phillips et al. 2014). Additionally, the RNA-dependent RNA
polymerase RRF-1 and the Zc3h12a-like ribonuclease RDE-8 localize to Mutator foci,
however their dependencies for localization were not known (Phillips et al. 2012; Tsai et
al. 2015). No proteins have yet been identified that are required for MUT-16 localization,
suggesting that MUT-16 may be the primary mediator of Mutator foci formation.
Mutator foci are considered hubs of siRNA amplification because mutations in mut-16
or any of the associated Mutator complex proteins (mut-2, mut-7, mut-14 smut-1, mut-15,
rde-2, rde-8, or rrf-1) result in a substantial loss of the RdRP-dependent secondary
38
siRNAs (Gu et al. 2009; Gent et al. 2010; Zhang et al. 2011; Phillips et al. 2012; Phillips
et al. 2014; Tsai et al. 2015). Furthermore, Mutator foci reside adjacent to another
ribonucleoprotein (RNP) granule, the P granule, which contains additional proteins
associated with RNA silencing and mRNA decay (Phillips et al. 2012). Notably, the
Argonaute protein WAGO-1, which binds secondary siRNAs synthesized in the Mutator
complex, localizes to and presumably silences mRNAs complementary to its bound
siRNAs in the P granule (Gu et al. 2009; Conine et al. 2010). It is unclear how mRNAs
targeted by primary Argonaute proteins get trafficked to the Mutator foci or how siRNAs
generated in the Mutator foci end up bound by WAGO-1 in the P granule. However, the
close juxtaposition between the P granule, the Mutator foci, and the nuclear pore
suggests a role for sorting RNAs and proteins between compartments during the multi-
step process of mRNA recognition and RNA silencing.
Little is known about how MUT-16 nucleates Mutator foci, but one clue comes from its
protein sequence. MUT-16 is highly enriched in disorder-promoting amino acids such as
glutamine (Q) and proline (P) (Phillips et al. 2012; van der Lee et al. 2014). Intrinsically
disordered regions (IDRs), which lack predicted structure, are often found in proteins
linked to the formation of RNA granules (i.e. the nucleolus, P bodies, stress granules, and
germ granules) (Wang et al. 2014; Nott et al. 2015; Elbaum-Garfinkle et al. 2015; Molliex
et al. 2015; Lin et al. 2015; Patel et al. 2015; Berry et al. 2015; Feric et al. 2016). Transient
interactions between IDR-containing proteins can be a driving force for RNA granule
formation through protein condensation and liquid-liquid phase separation, which occurs
when proteins and RNAs self-organize to form a distinct compartment with liquid-like
characteristics, but separate from the cytoplasm or nucleoplasm. This liquid droplet or
39
granule, sometimes referred to as membrane-less organelle, can readily exchange
components with the surrounding cytoplasm or nucleoplasm (Nott et al. 2015; Lin et al.
2015; Feric et al. 2016; Saha et al. 2016). Thus, liquid-liquid phase separation can
facilitate reactions by increasing the local concentration of proteins or RNA, as has been
proposed for the nucleolus (Feric et al. 2016). Alternatively, condensation can segregate
certain factors away from the cytoplasm, for example sequestration of mRNAs and
translational repressors into P bodies (Hubstenberger et al. 2017).
Here we demonstrate that the assembly of the Mutator complex and the formation of
Mutator foci are mediated by the intrinsically disordered protein MUT-16. MUT-16 acts as
a scaffold to recruit RRF-1 and other Mutator complex proteins, and it promotes foci
formation through its C-terminal region. We further show that Mutator foci depend on
weak hydrophobic interactions for their formation and form in a concentration and
temperature-dependent manner. Mutator foci also recover rapidly after photobleaching.
Thus, our data suggest that Mutator foci are phase-separated compartments with liquid-
like properties that associate with P granules and nuclear pores in the cytoplasm of germ
cells to promote RNA silencing.

RESULTS
MUT-16 and orthologs contain predicted intrinsically disordered regions
In previous work, we observed that MUT-16 was both Q/N-rich and P-rich,
predominantly in its C-terminal region and contains no other conserved domains (Phillips
et al. 2012). Because amino acid sequences containing low complexity regions, such as
Q/N-rich or P-rich regions, are often associated with disorder, we sought to identify
40
regions of intrinsic disorder within the MUT-16 protein sequence using IUPred (Dosztányi
et al. 2005a; Dosztányi et al. 2005b). MUT-16 has a short IDR near the N-terminus (first
~100 amino acids) and a much longer IDR comprising approximately 60% of the protein
(Fig 1A). In total, more than 70% of the protein is predicted to be unstructured. To
determine if the intrinsically disordered nature of MUT-16 is a conserved feature of this
protein, we used IUPred to predict IDR within the orthologs of MUT-16, in C. remanei, C.
briggsae, and C. japonica. Despite the relatively low sequence conservation between
orthologs, particularly in the C-terminal half of the protein (Phillips et al. 2012), the
conservation of IDRs was striking (Fig 1A). Like in C. elegans, the C. remanei, C.
briggsae, and C. japonica MUT-16 proteins have a short IDR near the N-terminus and
much longer IDR comprising the majority of the middle to C-terminal portions of the
proteins.
To determine if the high incidence of Q/N/P residues is distributed throughout the IDR
of MUT-16 or if it is restricted to a subset of the IDR, we counted the number of each of
these residues in a sliding window of 100 amino acids, starting at position one and shifting
10 residues at a time. We observed a prominent peak of glutamine which overlapped with
and was somewhat preceded by a prominent peak of proline (Fig 1B). These peaks
spanned only a subset of the C-terminal IDR. Like the conservation of IDR, the pattern of
enrichment of Q/N/P residues across MUT-16 orthologs in C. remanei, C. briggsae, and
C. japonica is strikingly similar and suggestive of a functional role for Q/N/P residues (S1
Fig).

41
Small deletions in mut-16 are RNAi defective
Because IDRs have been shown to promote phase separation of proteins into liquid-
like droplets, and MUT-16 forms foci in vivo, we sought to establish whether some or all
of the MUT-16 IDR was required for Mutator foci formation. We also sought to identify
regions of MUT-16 that directly recruit other Mutator complex proteins (Fig 1C). To this
end, we generated a series of small deletions of the MUT-16 protein using CRISPR
genome editing (Fig 1D-E). Each deletion removes between 61 and 141 residues (6-13%)
of MUT-16. For simplicity, we refer to the deletions as ΔA through ΔL. We initially planned
to make twelve deletions but due to technical constraints, the ΔH and ΔI deletions were
combined to make the single, slightly larger ΔH-I deletion. We additionally generated two
large deletions that remove most or all of the large IDR of MUT-16, which we will refer to
as ΔE-I or ΔE-K.
A strain containing mut-16(pk710), a null mutation in the mut-16 gene, is defective in
germline and somatic exogenous RNAi (Tabara et al. 1999; Zhang et al. 2011; Phillips et
al. 2012). To determine the severity of RNAi defects caused by each mut-16 deletion, we
tested the response of each mutant strain to dsRNA targeting the germline gene pos-1,
which causes embryonic lethality. We also tested their response to the somatic genes
nhr-23, which causes larval arrest, and lin-29, which causes the intestine and gonad of
the animal to rupture through the vulva at the larval to adult transition. Surprisingly, our
RNAi assays revealed that only mut-16 deletions ∆C, ∆F, ∆H-I, and ∆L have a significant
impact on somatic RNAi (Fig 2A-B). The remaining deletions had only mild or no RNAi
defect. In contrast, all deletions in mut-16 had defects in germline RNAi, though deletions
in ∆A and ∆G had more modest effects (Fig 2C). These data reveal that the majority of
the MUT-16 protein is necessary for robust germline RNAi. In contrast, some regions of
42
MUT-16 are dispensable for the response to at least some somatic RNAi clones,
suggesting that the soma may be more resilient to mild perturbations in MUT-16.

The C-terminal region of MUT-16 is critical for Mutator foci formation
Each of the mut-16 deletions were generated in the mut-16::mcherry::2xHA
background. To determine which of these regions is required to promote foci formation,
we examined the localization of mut-16 in each deletion background by live imaging (Fig
3A-E). Most deletions, including ΔA, ΔB, ΔD, ΔE, ΔF, ΔG, and ΔH-I, formed foci with
similar intensity to the control (full-length) strain (Fig 3A-B). The ΔC mutation still formed
Mutator foci, but had an overall reduced fluorescence (Fig 3A, 3C). It is unclear whether
the reduced fluorescence intensity observed in the ΔC mutation is due to reduced
expression or stability of MUT-16 specifically in the germline, but the overall MUT-16
protein levels are similar to full-length MUT-16 (Fig 3F). In contrast, ΔJ, ΔK, and ΔL
displayed severely disrupted foci, though the overall expression of cytoplasmic MUT-16
was not reduced (Fig 3A, 3D). We also examined the two large deletions, ΔE-I and ΔE-
K, which remove most of the IDR. Surprisingly, ΔE-I, which removes 387 amino acids
(~37% of the protein) and encompasses a substantial portion of the IDR, does not affect
MUT-16 localization, indicating that a large portion of the IDR is dispensable for foci
formation. In contrast, ΔE-K does disrupt MUT-16 localization, which is not unexpected
given that it includes J and K regions, which individually disrupted foci formation. All
deletions were expressed at similar or higher levels than the full-length strain (Fig 3F),
indicating that the reduced level of foci in ΔJ, ΔK, ΔL, and ΔE-K, is not due to lower protein
expression. Together, these data indicate that C-terminal region of MUT-16 (J, K, and L)
contains a region essential for foci formation.
43

Mutator proteins are each recruited by different regions of MUT-16
In previous work we examined the requirements for Mutator foci formation (Phillips et
al. 2012). In brief, MUT-16 is required for localization of MUT-2, MUT-7, RDE-2, MUT-14,
and MUT-15, all of which localize independently of one another except for MUT-7, which
requires RDE-2 for localization (Fig 1C). RRF-1 and RDE-8 have both been previously
shown to colocalize and co-immunoprecipitate with MUT-16 but whether they directly
interact with MUT-16 or interact via other Mutator complex proteins was unclear (Phillips
et al. 2012; Phillips et al. 2014; Tsai et al. 2015). We also suspected the Zc3h12a-like
ribonucleases NYN-1 and NYN-2 to be Mutator complex proteins because we had
identified them in an IP-mass spectrometry experiment with MUT-16 (S1 Table).
Additionally, Tsai et al. (2015) had identified them in a IP-mass spectrometry experiment
with RDE-8, as well as demonstrated that the nyn-1; nyn-2 double mutant displayed a
marked reduction in WAGO-class 22G-RNAs, a phenotype similar to that of mutations in
other Mutator complex components. To determine whether NYN-1 localizes to the
Mutator foci and to define the requirements for RRF-1, RDE-8, and NYN-1 localization,
we generated GFP::RRF-1, mCherry::RDE-8, and mCherry::NYN-1 using CRISPR. Each
protein forms distinct foci in germ cells and colocalizes with MUT-16, indicating that NYN-
1 is indeed a component of the Mutator foci (S2A Fig). Tagged RRF-1, RDE-8, and NYN-
1 strains were then crossed to strong loss-of-function mutations in other known Mutator
foci components, including mut-16, mut-2, rde-2, mut-14 smut-1, and mut-15. Of the
genes tested, only mut-16 was required for RRF-1 localization (S2B Fig). Localization of
RDE-8 and NYN-1 to Mutator foci requires both mut-16 and mut-15. To determine if NYN-
1 or RDE-8 were required for each other’s localization we further crossed mCherry::NYN-
44
1 to rde-8 mutants, and mCherry::RDE-8 to nyn-1; nyn-2 double mutants. mCherry::NYN-
1 was able to localize independently of RDE-8, whereas NYN-1 and NYN-2 were required
for RDE-8 localization (S2B Fig).
To identify regions of the MUT-16 protein that are required for recruitment of other
Mutator complex proteins we introduced the panel of mut-16 deletions into the MUT-
2::GFP, mCherry::RDE-8, mCherry::NYN-1, GFP::RRF-1, RDE-2::GFP, and MUT-
14::GFP strains. Each of these six lines had a wild-type RNAi response in the presence
of full-length mut-16 (S3A Fig) (Phillips et al. 2012). All deletion strains were generated
independently of one another and deletions of the same region in different strain
backgrounds behaved similarly to one another with respect to their effect on MUT-16
localization and ability to respond to germline and somatic RNAi (S3B Fig). MUT-2::GFP
foci were disrupted in the ΔB, ΔC, ΔJ, ΔK, and ΔL deletions (Fig 4A-D and S4 Fig),
however MUT-16 localization was also disrupted in ΔJ, ΔK, and ΔL (Figs 3A, 3D, 4D). To
determine whether MUT-2 and MUT-16 could still interact at the molecular level in each
deletion strain, we performed co-immunoprecipitation in each of the MUT-16 deletion
backgrounds. The interaction between MUT-2 and MUT-16 was only disrupted in the ΔB
and ΔC deletions, however, disruption of interactions between MUT-16 ΔC and other
Mutator complex proteins could be at least partially due to the reduced expression of
MUT-16 ΔC in the germline (S5 Fig). Despite the reduction in visible Mutator foci in ΔJ,
ΔK, and ΔL, MUT-2 still physically interacts with MUT-16 at levels similar to the full-length
control when these deletions are present (Fig 4E), indicating that ΔJ, ΔK, and ΔL regions
are not required for recruitment of MUT-2 to the Mutator complex. This result suggests
that when visible Mutator foci are disrupted in the ΔJ, ΔK, and ΔL mutants, many of the
45
Mutator complex proteins may still interact in diffuse cytoplasmic complexes. We also
observed that the MUT-16 protein is highly prone to degradation during the
immunoprecipitation procedure. While full-length MUT-16 could be observed in most
lanes, we also could detect multiple degradation products, with a prominent product of
~70kD in most lanes (S5 Fig). We did not observe substantial degradation with other
proteins we worked with and suspect the highly disordered nature of MUT-16 may leave
it more exposed to proteases during the immunoprecipitation procedure.
Similar to MUT-2 interaction, RDE-8, NYN-1, and MUT-14 depend on the ΔB and ΔC
regions of MUT-16 for localization to the Mutator foci (Fig 4A and S4 Fig). In contrast,
RDE-2 fails to localize to Mutator foci in the ΔH-I deletion, and RRF-1 is at least partially
disrupted in ΔB, ΔC, ΔF, and ΔG mutants (Fig 4A and S4 Fig). Because RRF-1 foci are
more difficult to detect than the other Mutator complex proteins, we also tested the
physical interaction between MUT-16 and RRF-1 by co-immunoprecipitation in each of
the MUT-16 deletion backgrounds. Only the ΔF mutant completely disrupted the
interaction between MUT-16 and RRF-1, though the amount of RRF-1
immunoprecipitated by the ΔB, ΔC, and ΔG mutants was modestly reduced relative to
full-length MUT-16 (Fig 4F). All together, these results suggest that different regions of
MUT-16 are important for recruiting each of the Mutator complex proteins. These regions
are separate and distinct from the C-terminal J, K, and L regions, which are important for
Mutator foci formation.

The C-terminal region of MUT-16 is sufficient for foci formation
The J, K, and L regions of MUT-16 are each necessary for robust MUT-16 foci
formation; to determine whether they are also sufficient for foci formation, we generated
46
a series of C-terminal fragments of MUT-16 fused to GFP and inserted them into the
genome using MosSCI (Fig 5A-B) (Frøkjaer-Jensen et al. 2008). Full-length MUT-16
forms foci throughout the germline with the brightest foci present in the mitotic region and
the leptotene/zygotene regions of the germline (Fig 5C) (Phillips et al. 2012). To
guarantee that any visible foci are not seeded by full-length, untagged MUT-16, C-
terminal MUT-16 fragments were introduced into a strain containing an early stop codon
in the mut-16 gene prior to imaging. All C-terminal GFP fusion constructs containing the
JKL region (mut-16
EFGHIJKL
::gfp, mut-16
GHIJKL
::gfp, mut-16
IJKL
::gfp, and mut-16
JKL
::gfp)
had visible MUT-16 foci, but as the fusion proteins became smaller (mut-16
IJKL
::gfp and
mut-16
JKL
::gfp), the foci became less intense and fewer in number (Fig 5D-G). mut-
16
KL
::gfp did not form foci, even in the presence of an endogenous, full-length copy of
MUT-16 and despite robust expression of this MUT-16 fragment (Fig 5B and 5H). These
data are consistent with the JKL regions being necessary and sufficient for foci formation,
but also indicate that additional regions of MUT-16 may help promote robust Mutator foci.

MUT-16 foci formation is concentration dependent
We previously observed that Mutator foci form in the germline but not in somatic cells
(Phillips et al. 2014). In the process of constructing an N-terminally GFP-3xFLAG-tagged
MUT-16, a strain construction intermediate was a mut-16 transcriptional reporter (mut-
16p::gfp) at the endogenous mut-16 locus (Dickinson et al. 2015). While imaging mut-
16p::gfp, we observed that, while GFP fluorescence is present throughout the animal, the
germline is distinctly brighter, suggesting that MUT-16 may be expressed at higher levels
in the germline compared to the somatic tissues (Fig 6A-B). Numerous studies have
shown that phase separation is a concentration-dependent process (Nott et al. 2015;
47
Elbaum-Garfinkle et al. 2015; Molliex et al. 2015; Lin et al. 2015; Weber et al. 2015). We
hypothesized that MUT-16 foci may form in germ cells but not in somatic cells because
MUT-16 germ cell protein levels are above the concentration threshold at which MUT-16
phase separates to form foci. To test this hypothesis, we generated myo-3p::mut-16::gfp,
which drives MUT-16 from a muscle-specific promoter and is expressed from a high-copy
extrachromosomal array. This transgenic strain, in which the MUT-16 protein is
overexpressed in muscle cells, has muscle-specific MUT-16::GFP foci (Fig 6C, top row).
The foci vary in size, some being substantially larger than endogenously expressed MUT-
16 foci. The co-expressed mCherry protein (myo-3::mCherry), which we also used as a
marker for muscle cells, does not form foci but rather is expressed diffusely in both the
cytoplasm and nucleus. Similarly, the GFP protein alone expressed under the myo-3
promoter (myo-3::gfp) and injected at the same molar concentration as myo-3p::mut-
16::gfp, is expressed diffusely in the cytoplasm and nucleus of muscle cells (Fig 6C,
bottom row). A control strain which expressed only myo-3p::mCherry in the presence of
mut-16::gfp expressed at the endogenous locus under its endogenous promoter (mut-
16p::mut-16::gfp) did not have foci in muscle cells (Fig 6C, middle row). Rather, diffuse
cytoplasmic expression of endogenous mut-16p::mut-16::gfp is visible in the muscle and
throughout the animal, indicating that over-expression of the mCherry protein in the
muscle does not drive MUT-16 into ectopic foci. To determine if the size or quantity of
ectopic MUT-16 foci change at different protein concentrations, we injected the myo-
3p::mut-16::gfp plasmid at 20 ng/ul, 5 ng/ul, 1 ng/ul or 0.25 ng/ul. While there was
variability between independent lines isolated from each set of injections, generally lines
from injections at the highest concentration (20 ng/ul) had larger and brighter foci than
48
lines from lower concentrations (5 ng/ul, 1 ng/ul, or 0.25 ng/ul), whereas some lines from
the lower concentration injections had no visible foci at all (Fig 6D). In some very high
expressing 20 ng/ul lines, we observed large condensates of MUT-16::GFP protein that
are no longer spherical (Fig 6D, top left). The presence of these larger structures could
indicate that at very high concentrations MUT-16 is behaving more solid or gel-like, which
has been seen previously for some IDR-containing proteins Lin et al. 2015). These data
indicate that overexpression of MUT-16 in muscle cells is sufficient for somatic foci
formation and suggest that MUT-16 can form foci in a concentration-dependent manner.
To determine whether the ectopic foci formed by overexpression of MUT-16 in the
muscle can recruit other Mutator proteins, we generated extrachomosomal arrays
overexpressing myo-3p::mut-16::gfp in strains carrying MosSCI lines expressing either
mut-15::mCherry or rde-2::mCherry. In a control strain expressing only myo-3p::mut-
16::gfp; there is very little mCherry signal overlapping the ectopic MUT-16 foci (Fig 6E,
top row). In contrast, in the strains expressing mut-15::mCherry or rde-2::mCherry at
endogenous levels, ectopic MUT-16 foci in the muscle recruit detectable levels of mut-
15::mCherry or rde-2::mCherry (Fig 6E, middle and bottom rows). Thus overexpression
of MUT-16 is sufficient to drive not just MUT-16, but also other Mutator complex proteins,
into ectopic Mutator foci.

Formation of Mutator foci depend on temperature and hydrophobic interactions
Multiple studies have demonstrated that liquid-like condensates, but not solid
aggregates, can be disrupted by aliphatic alcohols, such as 1,6-hexanediol (Updike et al.
2011; Kroschwald et al. 2015; Strom et al. 2017; Rog et al. 2017). These compounds
disrupt weak, hydrophobic interactions, and have previously been shown to alter the
49
permeability of the nuclear pore (Ribbeck et al. 2002). Addition of 5% 1,6-hexanediol to
C. elegans gonads resulted in severely disrupted MUT-16 foci (Fig 7A). Because a liquid-
like state requires weak molecular interactions whereas tighter interactions promote a
solid state (Hyman et al. 2014), these results support the hypothesis that Mutator foci
have liquid-like properties.
Phase-separated condensates can also be perturbed by changes in temperature. For
example, the DEAD-box helicase Ddx4, which is a component of germ granules, contains
disordered regions that condense upon exposure to low temperatures and dissolve when
returned to high temperatures (Nott et al. 2015). Other disordered proteins behave in the
opposite manner, condensing at high temperatures and dissolving at low temperatures
Jiang et al. 2015). To determine whether MUT-16 foci are temperature-sensitive, we
subjected MUT-16::GFP animals to heat stress by placing them at 30°C for 6 hours. We
returned the animals to room temperature (~21°C), and strikingly, the MUT-16 foci were
no longer visible in the germ cells (Fig 7B and S6). However, within 15 minutes, some
animals have already reformed MUT-16 foci (Fig 7B and S6). Over the full time-course of
60 minutes at room temperature, many heat-shocked animals displayed no discernable
differences in foci presence and intensity from animals raised at permissive temperatures
(Fig 7B, S6 Fig, and S1 Movie). These data indicate that MUT-16 foci can change in
response to environmental conditions, such as temperature.

Mutator foci recover after photobleaching
Germ granule proteins, such as PGL-1, PGL-3, and LAF-1 in C. elegans P granules,
recover rapidly after photobleaching indicating that the internal components of the granule
50
can rearrange, similar to molecules in a liquid state (Brangwynne et al. 2009; Sheth et al.
2010; Elbaum-Garfinkle et al. 2015; Saha et al. 2016). To determine if MUT-16 foci are
similarly liquid-like, we performed fluorescence recovery after photobleaching (FRAP)
experiments. Due to the small size of Mutator foci (<500 nm), we chose to bleach entire
foci and measure the recovery of MUT-16 to the foci from the surrounding cytoplasm.
MUT-16::GFP foci recovered rapidly after photobleaching, t1/2 = 7.2 ± 1.0 seconds (SEM,
n = 5). Recovery only reaches ~35% of pre-bleached intensity (Fig 7C-D and S1 Movie),
indicating that there may be both a mobile fraction of MUT-16 that can exchange quickly
between Mutator foci and the cytoplasm, and an immobile fraction that exchanges very
slowly or not at all. Incomplete recovery of fluorescence has previously been observed
for in vivo FRAP of P granules and of the adjacent and recently discovered Z granules
(Brangwynne et al. 2009; Sheth et al. 2010; Wan et al. 2018). Fluorescence recovery
data, together with the dependence of Mutator foci on concentration, temperature, and
hydrophobic interactions, suggest that Mutator foci have properties of a phase-separated
condensate.

DISCUSSION
Functional subdivision of MUT-16
MUT-16 function can be subdivided across different regions of the protein (Fig 7E).
The largest structured region (specifically the B-C region) of MUT-16 recruits multiple
proteins, including the nucleotidyl transferase MUT-2, the DEAD-box RNA helicase MUT-
14, and the protein of unknown function MUT-15, whose localization requirements we can
infer based on the localization of the Zc3h12a-like ribonucleases RDE-8 and NYN-1 (Figs
51
4A, 4C, and S2B). MUT-2, MUT-14, and MUT-15 likely interact directly with MUT-16, as
we can robustly detect these interactions by immunoprecipitation and no other unknown
proteins have been identified as part of this complex by IP-mass spectrometry
experiments (Figs 4, S5, and S1 Table) (Phillips et al. 2012). Since MUT-15 recruits NYN-
1, NYN-2 and RDE-8, at least six Mutator complex proteins localize to Mutator foci
through the B-C region of MUT-16 (Figs S2B and 7E). It remains to be determined
whether a single MUT-16 protein can interact with all of these proteins concurrently.
Interestingly, while we initially hypothesized that the IDR would be important for
promoting Mutator foci formation, we found regions of disorder also function to recruit
proteins to the Mutator complex. In particular, region F recruits the RdRP protein RRF-1
and the H-I region recruits the exonuclease MUT-7 through its interaction with RDE-2.
The regions that directly recruit Mutator complex proteins correlate with regions required
for robust RNAi in somatic tissues (Fig 2). Specifically, mut-16∆C, ∆F, and ∆H-I have
somatic RNAi defects nearly as severe as a mut-16 null allele. mut-16∆L also has similarly
severe somatic RNAi, however unlike ∆C, ∆F, and ∆H-I, deletion of the L region does not
disrupt the interaction of MUT-16 with any other Mutator complex proteins of which we
are aware. Since we cannot detect Mutator foci in the soma, we do not yet know how the
loss of the L region affects Mutator complex formation or function in somatic cells. In
addition, it is unclear why the remaining regions of MUT-16 are required for germline but
not somatic RNAi. It is possible that they recruit yet uncharacterized germline-specific
RNAi factors, or may facilitate currently unexamined interactions between primary and
secondary Argonaute proteins and the Mutator complex. Overall, this data suggests that
at least some of the most severe defects in somatic RNAi that occur after deleting portions
52
of the mut-16 gene stem from loss of recruitment of specific proteins to the Mutator
complex.
Our fusion of C-terminal fragments of MUT-16 to GFP, demonstrates that a minimal
region comprised of amino acids 773-1050 (JKL region) is sufficient for Mutator foci
formation (Fig 5). This region is ~70% disordered, but contains the structured L region
that is also key to the formation of Mutator foci. We did observe that including additional
regions of the MUT-16 protein, specifically a larger portion of the disordered region, can
promote increased size and number of Mutator foci. Interestingly, the regions with the
highest glutamine and proline content are centered on the H-I and J regions of MUT-16,
only partially overlapping this region critical for Mutator foci formation. This finding
suggests that isolated stretches of disordered amino acids are not sufficient to mediate
phase separation, but require the neighboring protein environment to promote Mutator
foci formation.

MUT-16 foci have properties of a phase-separated condensate
Only recently has it become clear that many biological processes involve intracellular
phase transitions, from spindle assembly to heterochromatin formation (Jiang et al. 2015;
Strom et al. 2017; Larson et al. 2015). Proteins that are heterogeneous in conformation,
such as proteins containing IDR, typically drive phase separation by providing a flexible
platform for non-covalent interactions with nearby disordered proteins and/or ribonucleic
acids. Other conditions such as protein or RNA concentration, post-translational
modification, or changes in environmental conditions such as salt concentration or
temperature can modulate these transitions (Brangwynne et al. 2015). Interestingly, we
have been unable to co-immunoprecipitate MUT-16 with itself, which, while not
53
conclusive, suggests that MUT-16 does not form strong intermolecular interactions and
rather promotes Mutator foci formation via many weak or transient interactions.
Based on our studies of MUT-16, we propose that Mutator foci are phase-separated
membrane-less organelles with liquid-like properties rather than a solid or aggregated
structure, with five lines of evidence supporting this hypothesis. First, Mutator foci are
roughly spherical in shape, as would be expected of a liquid compartment due to the
cohesive forces of the surface layer (i.e surface tension). Second, Mutator foci dissolve
in the aliphatic alcohol, 1,6-hexanediol, which disrupts only weak, hydrophobic
interactions but not solids. Third, MUT-16::GFP foci recover rapidly, though incompletely,
after photobleaching, indicating that a mobile fraction of MUT-16 can exchange freely
between the Mutator foci and the cytoplasm. Fourth, Mutator foci only assemble when
concentrations rise above a certain threshold, which is a hallmark of phase-separated
condensates. And fifth, Mutator foci formation can be modulated by changes in
temperature – increased entropy at higher temperatures can counteract phase separation
and promote mixing of the condensed and bulk phases. The latter two points indicate that
Mutator foci may be sensitive to intracellular and extracellular conditions, which could
allow foci formation to be fine-tuned based on cell type or environmental conditions.
A liquid-like nature of Mutator foci could allow for some proteins and RNA to freely
exchange in and out of the compartment, while still maintaining a high concentration of
factors required for small RNA amplification. Furthermore, the close juxtaposition of
Mutator foci with nuclear pores and P granules, which also have liquid-like properties,
suggests a model of RNA silencing where adjacent membrane-less organelles can
exchange proteins and RNAs, but have intrinsic properties that make them immiscible
54
with one another. A similarly close juxtaposition has been observed between P bodies
and stress granules, and these types of interactions may reflect differences in surface
tension of the two types of condensates (Kedersha et al. 2005; Shin and Brangwynne et
al. 2017). Further characterization both in vivo and in vitro will be necessary to fully
elucidate how Mutator foci form and interact with neighboring membrane-less organelles.

Properties of the Mutator complex in the soma vs. germline
Why are Mutator foci present in the germline but not the soma? The answer may lie
in the unique biophysical properties of MUT-16, as well as in the concentration of and
local environment surrounding MUT-16 in the germline. By concentrating Mutator
complex proteins to the cytoplasmic side of nuclear pores, Mutator foci may be poised to
capture deleterious RNAs during nascent RNA export. It is unclear why a similar
mechanism is not also in place in somatic cells, but perhaps such a robust RNA silencing
mechanism is not necessary in the soma as these cells will not be passed to the next
generation.
Mutator foci in the germline are exclusively associated with the nuclear periphery, until
the germ cells become oocytes and the P granules detach from the nuclei and become
cytoplasmic. Mutator foci and some nuclear pore components move with the P granules
into the cytoplasm (Pitt et al. 2000; Sheth et al. 2010; Phillips et al. 2012). We do not yet
know whether Mutator foci can interact directly with nuclear pores or if this interaction is
mediated by the P granule or another unknown factor, however knockdown of P granule
components does not disrupt the perinuclear localization of Mutator foci (Phillips et al.
2012). Interestingly, while some of the MUT-16 foci in myo-3p::mut-16::gfp are adjacent
to the nuclear periphery, most are not (Fig 6C). It is unclear whether the lack of perinuclear
55
association in the muscle cells is due to the lack of a specific anchoring factor present
only in germ cells, or perhaps the anchoring factor (for example, nuclear pores) is present
but limited in supply relative to the substantial overexpression of MUT-16.
In conclusion, our results demonstrate that MUT-16 serves as a scaffold for assembly
of Mutator foci and suggest that these foci form through liquid-liquid phase separation.
The association of Mutator foci with P granules and nuclear pores suggests a model
whereby newly transcribed mRNAs pass through nuclear pores into the P granule, where
mRNAs are marked for silencing and targeted to the neighboring Mutator foci.
Subsequently, siRNAs synthesized in the Mutator foci and perhaps secondary Argonaute
proteins can transit back into the P granules to target additional complementary mRNAs
for silencing. Thus, we propose that RNA silencing in germ cells may depend on RNA
transit through phase-separated liquid compartments that concentrate RNA silencing
enzymes.

METHODS
Strains
The C. elegans wild-type strain is N2. Worms were cultured at 20°C according to
standard conditions unless otherwise stated (Brenner 1974). Strains used include WM30
(mut-2(ne298) I), NL3531(rde-2(pk1657) I), NL1810 (mut-16(pk710) I), FX04844 (nyn-
2(tm4844) I), NL1820 (mut-7(pk720) III), HT1593 (unc-119(ed3) III), EG4322 (ttTi5605 II;
unc-119(ed9) III), GE24 (pha-1(e2123) III), FX05004 (nyn-1(tm5004) IV), FX02252 (rde-
8 (tm2252) IV), GR1948 (mut-14(mg464) smut-1(tm1301) V), and GR1747(mut-
56
15(tm1358) V). All mutants were outcrossed at least 4x prior to any analysis. New strains
made for this project are listed in the S2 Table associated with the original publication.

Plasmid and strain construction
All GFP and mCherry constructs were designed for insertion at the endogenous loci
by CRISPR genome editing except the C-terminal fragments of mut-16 fused to GFP (Fig
5), which were integrated by Mos-mediated single-copy transgene insertion (MosSCI)
(Frøkjaer-Jensen et al. 2008). For all CRISPR insertions of GFP or mCherry, we
generated homologous repair templates using the primers listed in S3 Table associated
with the original publication. To create the mut-16::gfp::3xFLAG repair template, we first
generated a GFP with C-terminal 3xFLAG tag and internal Floxed Cbr-unc-119(+), and
inserted into the Kanamycin-resistant backbone of pDONR221 by isothermal assembly
(Gibson et al. 2009). This cassette was then flanked with ~1.5-2kb of sequence from
either side of the mut-16 stop codon, again by isothermal assembly. A similar design was
used to create all mCherry repair templates. We first generated a
2xHA::mCherry(w/internal Floxed Cbr-unc-119(+)]::2xHA construct by cloning
mCherry(w/internal Floxed Cbr-unc-119(+)] into XhoI/SpeI-digested pBluescript SK(-).
The mCherry[w/internal Floxed Cbr-unc-119(+)] was a gift from the lab of Jeremy Nance
(NYU). BamHI and BglII sites were engineered into the mCherry construct just following
the ATG or just preceding the stop codon, respectively. This construct was sequentially
digested with BglII, then BamHI; each digest was followed by ligation of a 2xHA oligo.
This construct was then used as a PCR template to generate mCherry::2xHA[w/internal
Floxed Cbr-unc-119(+)] or 2xHA::mCherry[w/internal Floxed Cbr-unc-119(+)], and
assembled with ~1kb of sequence from either side of the mut-16 stop codon, or the rde-
57
8 and nyn-1 start codons, respectively. mut-2::gfp::3xFLAG, rde-2::gfp::3xFLAG,
gfp::3xFLAG::rrf-1, and gfp::3xFLAG::mut-16 (for mut-16p::gfp) repair templates were
assembled into pDD282 (GFP/3xFLAG with self-excising cassette, Addgene #66823)
according to published protocols (Dickinson et al. 2015). To protect the repair template
from cleavage, we introduced silent mutations at the site of guide RNA targeting by
incorporating these mutations into one of the homology arm primers or, if necessary, by
performing site-directed mutagenesis (Dickinson et al. 2013).
The mut-16 guide RNA was cloned into PU6::unc-119_sgRNA (Addgene #46169) by
site-directed mutagenesis (Friedland et al. 2013). All other guide RNA plasmids were
generated by ligating oligos containing the guide RNA sequence into BsaI-digested
pRB1017 (Addgene #59936) (Arribere et al. 2014). Guide RNA sequences are provided
in the S3 Table of the original publication.
C-terminal fragments of mut-16 fused to GFP were cloned into targeting vectors for
MosSCI. The endogenous promoter and 3’UTR were amplified, along with either full-
length coding sequence or C-terminal fragments of the coding sequence (S3 Table).
These amplicons were inserted along with GFP by isothermal assembly into SpeI-
digested pCFJ151 (Addgene #19330) (Frøkjaer-Jensen et al. 2008; Gibson et al. 2009).
A similar strategy was used to generate myo-3p::mut-16::gfp, except with the mut-16
promoter replaced by the myo-3 promoter.
CRISPR injections were performed according to published protocols (Dickinson et al.
2013; Dickinson et al. 2015; Ward 2015). GFP/mCherry CRISPR injection mixes included
10-25 ng/μl repair template, 50 ng/μl guide RNA, 50 ng/μl eft-3p::cas9-SV40_NLS::tbb-2
3'UTR (Addgene # 46168), 2.5-10 ng/μl GFP or mCherry co-injection markers, and 10
58
ng/μl hsp-16.1::peel-1 negative selection (pMA122, Addgene #34873). mut-
16::gfp::3xFLAG and all mCherry constructs were injected into HT1593 (unc-119(ed3)
III). Floxed Cbr-unc-119(+) cassettes were later excised using eft-3p::Cre (pDD104,
Addgene #47551) (Dickinson et al. 2013). mut-2::gfp::3xFLAG, rde-2::gfp::3xFLAG, and
gfp::3xFLAG::rrf-1 were injected into the wild-type strain. mut-16 deletion injection mixes
included 50 ng/μl oligo repair template, 25 ng/μl each of two mut-16 guide RNAs, 50 ng/μl
pha-1 repair oligo and 50 ng/μl eft-3p:Cas9 + pha-1 guide RNA (pJW1285, Addgene
#61252). Mixtures were injected into pha-1(e2123) mutant animals already carrying mut-
16::gfp::3xFLAG or mut-16::mCherry::2xHA and other fluorescently-tagged Mutator
complex proteins and mut-16 deletions were identified by PCR according to pha-1 co-
conversion protocol (Ward et al. 2015).
For MosSCI injections, we integrated transgenes into the ttTi5605 MosI site in strain
EG4322 (Ch. II) following a published MosSCI protocol (Frøkaer-Jensen et al. 2008).
Injection mixes contained 50 ng/μl MosSCI-targeting vector, 50 ng/μl eft-3p::Mos1
transposase (pCFJ601, Addgene #34874), 10 ng/μl rab-3p::mCherry (pGH8, Addgene
#19359), 2.5 ng/μl myo-2p::mCherry (pCFJ90, Addgene #19327), 5 ng/μl myo-
3p::mCherry (pCFJ104, Addgene #19328), and 10 ng/μl hsp-16.1::peel-1 negative
selection (pMA122, Addgene #34873). Extra-chromosomal arrays were generated as
follows: 20ng/ul myo-3p::mut-16::gfp, 5 ng/μl myo-3p::mCherry (pCFJ104) and 70 ng/ul
pBluescript injected into HT1593 - unc-119(ed3) for cmpEx76 and cmpEx89; 5 ng/ul myo-
3p::mut-16::gfp, 5 ng/μl myo-3p::mCherry (pCFJ104) and 70 ng/ul pBluescript injected
into HT1593 - unc-119(ed3) for cmpEx90; 1 ng/ul myo-3p::mut-16::gfp, 5 ng/μl myo-
3p::mCherry (pCFJ104) and 70 ng/ul pBluescript injected into HT1593 - unc-119(ed3) for
59
cmpEx91; 0.25 ng/ul myo-3p::mut-16::gfp, 5 ng/μl myo-3p::mCherry (pCFJ104) and 70
ng/ul pBluescript injected into HT1593 - unc-119(ed3) for cmpEx92; 5 ng/μl myo-
3p::mCherry (pCFJ104) and 70 ng/ul pBluescript injected into the mut-16(cmp3[mut-
16::gfp::3xFLAG]) strain for cmpEx79; 7.5 ng/ul myo-3p::gfp (pPD118.20) and 70 ng/ul
pBluescript into the wild-type strain for cmpEx88; and 20ng/ul myo-3p::mut-16::gfp and
70 ng/ul pBluescript injected into HT1593 - unc-119(ed3), mut-15::mCherry, and rde-
2::mCherry strains for cmpEx76, cmpEx86, and cmpEx87.

Antibody staining and imaging
C. elegans were imaged in M9 buffer with sodium azide to prevent movement. In some
cases, to obtain higher quality images, animals were dissected prior to mounting. For
scoring presence or absence of foci, deletions were blinded and scored by multiple
individuals. A minimum of 10 animals were scored for each strain. Imaging was performed
on a DeltaVision Elite microscope (GE Healthcare) using a 60x N.A. 1.42 oil-immersion
objective. When data stacks were collected, three-dimensional images are presented as
maximum intensity projections. Images were pseudocolored using the SoftWoRx
package or Adobe Photoshop.
For quantification of somatic and germline fluorescence intensity in the mut-16p::gfp
strain, 31 age-matched L4 animals were imaged using identical microscope settings. The
somatic and germline tissue was traced in ImageJ and mean gray value was calculated
for each region. 1,6-hexanediol treatment was performed by dissecting adult MUT-
16::GFP animals in M9 buffer with or without the addition of 5% 1,6-hexanediol and
imaged immediately following dissection. For heat stress experiment, plates of L4 stage
MUT-16::GFP animals were wrapped in Parafilm and placed in a 30°C incubator for 6
60
hours. We then removed the plates from the incubator, immediately transferred the
animals to slides, and began imaging. No more than 5 minutes elapsed between removal
from 30°C and collection of the first time point (Fig 7B, 0 min). Images were captured
every 3 minutes for 60 minutes at room temperature (~21°C).
Fluorescence recovery after photobleaching (FRAP) was performed on a Leica SP8
Falcon laser scanning confocal microscope using a 63x N.A. 1.4 oil-immersion objective.
Two images were acquired prior to bleaching, followed by bleaching, then 10 images at
0.44 second intervals and 10 additional images at 10 second intervals. Data was analyzed
using the Leica Application Suite X software.

Immunoprecipitation and western blots
Immunoprecipitations were performed as previously described (Phillips et al. 2012).
~40,000 synchronized adult C. elegans (~68 h at 20°C after L1 arrest) were collected,
frozen in liquid nitrogen, and homogenized using a mortar and pestle. After further dilution
into lysis buffer (1:10 packed worms:buffer), insoluble particulate was removed by
centrifugation and a sample was taken as “input.” The remaining lysate was used for the
immunoprecipitation. HA-tagged proteins were immunoprecipitated using anti-HA affinity
matrix (ThermoFisher 26181).
For Western blots, proteins were resolved on 4-12% Bis-Tris polyacrylamide gels
(ThermoFisher), transferred to nitrocellulose membranes, and probed with anti-HA
1:1,000 (Roche 12013819001), anti-FLAG 1:1,000 [M2 clone] (Sigma-Aldrich F1804),
anti-actin 1:10,000 (Abcam ab3280), or anti-GFP 1:100 (Riken BRC JFP-J5) (Hayashi
and Shirao 1999). Secondary HRP antibodies were purchased from ThermoFisher.
61
We observed that MUT-16 was often substantially degraded during the
immunoprecipitation procedure. We attempted to remedy this problem by reducing the
time samples spent in lysis buffer, shortening the immunoprecipitation step, and increase
the amount of protease inhibitor. These steps resulted in little improvement of full-length
MUT-16 yield in both input and IP samples. We also noted that the MUT-16 degradation
product was present in full-length and all MUT-16 deletion mutants except for ∆H-I. Given
that MUT-16 is tagged at the C-terminus, the deletions preceding ∆H-I all form
degradation products of the same size, and the deletions following ∆H-I all vary in size in
proportion to the relative deletion sizes, we suspect that the degradation site in MUT-16
is somewhere in the ∆H-I region (S5 Fig).
For mass spectrometry experiments, immunoprecipitation was performed as
described above, starting with ~500,000 synchronized adult C. elegans (~68 h at 20°C
after L1 arrest) for each sample. GFP and FLAG immunoprecipitation was performed
using anti-GFP affinity matrix [RQ2 clone] (MBL International D153-8) and anti-FLAG
affinity matrix [M2 clone] (Sigma-Aldrich A2220). After immunoprecipitation, samples
were precipitated using the ProteoExtract Protein Precipitation Kit (EMD Millipore
539180) and submitted to the Taplin Mass Spectrometry facility at Harvard Medical
School for protein identification.

RNAi assays
For RNAi assays, L1 animals were fed E. coli expressing dsRNA against pos-1, lin-
29, or nhr-23 (Kamath et al. 2003a). For pos-1, animals were scored 4 days later for
hatching of the F1 embryos. For lin-29 or nhr-23, animals were scored 2–3 d later for
vulval bursting or larval arrest, respectively. To quantify intermediate phenotypes of nhr-
62
23 and lin-29 RNAi, we established two subcategories to distinguish between partial RNAi
defects (animals are smaller than wild-type with few or no eggs as adults, and for lin-19,
often have a protruding vulva) and total RNAi defects (animals are phenotypically wild-
type).

ACKNOWLEDGMENTS
We would like to thank Taiowa Montgomery and members of the Phillips lab for
discussions and feedback on the manuscript, and Cosimo Arnesano, Anthony Fernandez,
Fabien Pinaud, and Scott Fraser for help with FRAP experiments. Some strains were
provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs
(P40 OD010440), and Shohei Mitani of the Japanese National BioResource Project.

63
CHAPTER 1 FIGURES

Figure 1. MUT-16 and orthologs contain a high degree of predicted disorder.
(A) Graph comparing disorder tendency of C. elegans MUT-16 (isoform b, WP:CE40347) in red
to orthologs in C. remanei (RP:RP48608) in light gray, C. briggsae (BP:CBP44329) in medium
gray, and C. japonica (JA:JA63728) in dark gray using IUPRED and long disorder parameters
(http://iupred.enzim.hu/). Scores above 0.5 indicate disorder. Residue positions correspond to the
C. elegans MUT-16 sequence, but all proteins are similar in length. (B) Q, N, and P residues in
C. elegans MUT-16 were counted in amino acid 100-mers, starting at position one, shifting 10
residues at a time, and displayed as stacked columns. Indicated residue positions are the mid-
point of the 100-mer. (C) Schematic of known dependencies of Mutator foci formation. The
requirements for localization of RRF-1, RDE-8, NYN-1, NYN-2, and SMUT-1 were not known prior
to this work. (D, E) Diagram (D) and table (E) of MUT-16 deletions generated by CRISPR. Bars
in (D) are drawn to scale relative to residue positions in (A,B).

64

Figure 2. Susceptibility of mut-16 deletion worms to somatic and germline RNAi.
(A-C) mut-16 deletion worms were assayed for their ability to respond to somatic (nhr-23 or lin-
29) or germline (pos-1) RNAi. For nhr-23 RNAi (A) and lin-29 RNAi (B), P0 animals were scored
as having intermediate RNAi defects (gray bars), or fully penetrant RNAi defects (red bars). For
pos-1 RNAi (C), the F1 eggs and hatched larvae were counted to calculate % viable progeny from
treated P0 animals. Weighted means and standard deviations were calculated from three
independent RNAi trials of n=~20 for nhr-23 and lin-29, and n=~90 F1 eggs from 4 P0 adults for
pos-1.

65

Figure 3. The C-terminal region of MUT-16 is necessary for Mutator foci formation.
(A) Table indicates whether MUT-16 foci are present or absent in each mut-16 deletion strain.
Yes indicates foci present in the majority of animals, No indicates foci absent or severely
disrupted, and Weak (for ∆C) indicates an intermediate phenotype where the fluorescence
intensity of cytoplasmic MUT-16 appeared reduced relative to the other deletion lines. (B-E) Live
imaging of MUT-16::mCherry expression and localization for control strain (B) or when ∆C (C),
∆L (D), or ∆E-I (E) deletions have been introduced into the mut-16::mCherry strain. Scale bars,
5µm. (F) MUT-16 western blot to assess protein levels in full-length and mut-16 deletion strains.
Expected sizes for MUT-16::mCherry::2xHA are 148 kD (full-length), 139 kD (∆A), 137 kD (∆B),
134 kD (∆C), 138 kD (∆D), 137 kD (∆E), 138 kD (∆F), 141 kD (∆G), 132 kD (∆H-I), 135 kD (∆J),
141 kD (∆K), 138 kD (∆L), 105 kD (∆E-I), and 85 kD (∆E-K). Approximately 200 synchronous adult
animals were loaded per lane and actin was used as a loading control.

66

Figure 4. Distinct regions of MUT-16 recruit each of the other Mutator proteins.
(A) Table indicates whether mut-16 deletions disrupt MUT-2, RDE-8, NYN-1, MUT-14, RRF-1,
and RDE-2 foci. Yes indicates foci present in the majority of animals, No indicates foci absent or
severely disrupted, and ND indicates that strain was not constructed or scored. (B-D) MUT-
16::mCherry and MUT-2::GFP expression and localization for control strain (B) or when ∆C (C)
or ∆K (D) deletions have been introduced into the mut-16::mCherry strain. Scale bars, 5 µm. (E)
Immunoprecipitation and western blot of MUT-16::mCherry::2xHA (expected sizes between 135
– 141 kD for MUT-16 deletions and 148 kD for MUT-16 full length) and MUT-2::GFP::3xFLAG (83
kD). Left panels are total lysate from strains indicated above, and right panels are following HA
immunoprecipitation. (F) Immunoprecipitation and western blot of MUT-16::mCherry::2xHA
(expected sizes between 132 – 142 kD for MUT-16 deletions and 148 kD for MUT-16 full length)
and GFP::3xFLAG::RRF-1 (219 kD). Top two panels are total lysate from strains indicated above,
and bottom two panels are following HA immunoprecipitation. The equivalent of ~0.5% of starting
material for the input fractions and ~20% of starting material for the IP fractions were loaded onto
the gels.

67

Figure 5. The C-terminal region of MUT-16 is sufficient for foci formation.
(A) Diagram and amino acid coordinates of C-terminal regions of MUT-16 fused to GFP. (B)
Western blot of full-length and C-terminal fragments of MUT-16::GFP probed with anti-GFP and
anti-actin. A non-specific band (marked by asterisk) runs at a similar size to the full-length MUT-
16::GFP. (C) Live imaging of full-length MUT-16::GFP, which forms foci throughout the germline.
(D-G) C-terminal fragments of MUT-16::GFP form foci in the germline if they contain the minimal
J, K, and L region (amino acids 773-1050) of the protein, but the number and size of foci increase
with larger protein fragments. (H) The KL region (amino acids 885-1050) of MUT-16 fused to GFP
is not sufficient for foci anywhere in the germline. All images are from the transition zone
(leptotene/zygotene) region of the germline. Scale bars, 5 µm.

68

Figure 6. Concentration dependence of Mutator foci.
(A) GFP expression in negative control (wild-type) and mut-16p::gfp L4 stage animals. The same
exposure time was used for both images. Scale bar, 50 µm. (B) Quantification of GFP
fluorescence in the soma and the germline of mut-16p::gfp animals. Fluorescence intensity was
calculated using ImageJ by tracing the germline and soma of 31 L4 stage animals and measuring
the mean gray value. P values were calculated using a paired t-test (*** indicates p<0.001). (C)
Top row: MUT-16::GFP overexpressed from the myo-3 promoter forms foci in muscle cells
marked with muscle-specific mCherry. Yellow arrow indicates the location of the nucleus. Middle
row: Endogenously expressed MUT-16::GFP in muscle cells marked with muscle-specific
mCherry does not form foci. White arrows point to MUT-16 foci in the neighboring germ cells.
Bottom row: Overexpression of the GFP protein alone in muscle cells does not form foci. Scale
bars, 10 µm. (D) MUT-16::GFP in muscle cells labeled with myo-3::mCherry, where myo-3p::mut-
16::gfp was injected at the indicated concentrations. Microscope exposure settings were changed
as follows to account for variation in brightness between lines - top left: 0.02 sec exposure with
neutral density filter at 10% transmittance; top right: 0.2 sec exposure; bottom left: 0.2 sec
exposure; bottom right: 0.4 sec exposure. Scale bars, 10 µm. (E) Top row: MUT-16::GFP
overexpressed from the myo-3 promoter forms foci. No mCherry proteins are expressed in this
strain. Middle and bottom rows: Overexpression of MUT-16::GFP in muscle cells recruits MUT-
15::mCherry (middle row) and RDE-2::mCherry (bottom row). Scale bars, 5 µm.
69

Figure 7. Mutator foci have liquid-like properties.
(A) MUT-16::GFP are present in buffer control, but disappear in 5% 1,6-hexanediol. Scale bar, 5
µm. (B) Images were collected at the indicated time points after MUT-16::GFP animals were heat-
shocked for 6 hours at 30°C and then returned to room temperature (~21°C). MUT-16 foci are
absent at the first time point (0 min), but begin to reform after ~15 min and look similar to untreated
MUT-16 foci by 30-60 min. Scale bar, 5 µm. (C) Time-lapse images show fluorescence recovery
of MUT-16::GFP after a single Mutator focus is photobleached at 2.0 sec. (D) FRAP recovery
curves for MUT-16::GFP. Raw data (gray line) is reported as a mean +/- SD (n = 5 granules).
Data was fitted to a single exponential recovery curve (red line). (E) Schematic of MUT-16,
indicating regions required for interaction with other Mutator complex proteins and for Mutator foci
formation. Shading along the length of the MUT-16 protein represents disorder tendency
calculated using IUPred (see Fig 1A).

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Figure S1. Distribution of Q, N, and P residues in MUT-16 orthologs.
The frequency of Q, N, and P residues were analyzed in C. elegans MUT-16 (WP:CE40347), C.
briggsae MUT-16 (BP:CBP44329), C. remanei MUT-16 (RP:RP48608), and C. japonica MUT-16
(JA:JA63728) proteins. Residues were counted in amino acid 100-mers, starting at position one,
shifting 10 residues at a time, and displayed as stacked columns. Indicated residue positions are
the mid-point of the 100-mer.
71

Figure S2. Requirements for RRF-1, NYN-1, and RDE-8 localization to Mutator
foci.
(A) GFP::RRF-1, mCherry::NYN-1, and mCherry::RDE-8 each colocalize with MUT-16. (B)
GFP::RRF-1, mCherry::NYN-1, and mCherry::RDE-8 were introduced into each of the indicated
mutant backgrounds. RRF-1 foci were disrupted in mut-16 mutants, NYN-1 were disrupted in mut-
16 and mut-15 mutants, and RDE-8 foci were disrupted in mut-16, mut-15, and nyn-1; nyn-2
double mutants. All images are from the transition zone (leptotene/zygotene) region of the
germline. Scale bars, 5 µm.
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Figure S3. Response of GFP- and mCherry-tagged strains to somatic and
germline RNAi.
(A) Deletion strain controls were assessed for RNAi response to somatic (nhr-23 and lin-29) or
germline (pos-1) RNAi. For somatic RNAi, P0 animals were categorized as having fully penetrant
(red bars) or intermediate RNAi defects (gray bars). Fully penetrant RNAi defects were larval
arrest for nhr-23, vulval burst for lin-29, and non-hatching F1 for pos-1. Intermediate effects were
categorized as non-thriving, slow moving adults with eggs for nhr-23 and as adults with protruding
vulvas for lin-29. For pos-1, the F1 eggs and hatched larvae were counted to calculate % viable
progeny from treated P0 animals. Weighted means and standard deviations were calculated from
three independent RNAi trials of n=~20 for nhr-23 and lin-29, and n=~140 F1 eggs from 4 P0
adults for pos-1. (B) Deletion strains were assessed for RNAi response and categorized as having
either fully penetrant RNAi defects (Rde, red boxes), intermediate RNAi defects (Weak, pink
boxes), or wild-type response (WT, white boxes). ND indicates that strain was not constructed or
scored. For nhr-23, fully penetrant RNAi defects are strains where >65% of animals are non-
arrested and healthy, intermediate RNAi defects are strains where >35% are non-arrested (either
healthy or sick), and wild-type response are strains where ≤35% are non-arrested (either healthy
or sick). For lin-29, fully penetrant RNAi defects are strains where >70% of animals were scored
as non-burst, intermediate RNAi defects are strains where 35%-70% of animals were scored as
non-burst, and wild-type response are strains where <35% are non-burst. For pos-1, fully
penetrant RNAi defects are strains where >75% F1s are viable, intermediate RNAi defects are
strains where 10-75% F1 are viable, and wild-type response are strains <10% F1 are viable.
73

74

Figure S4. Distinct MUT-16 regions are required for the localization of the other
Mutator proteins.
Live imaging of MUT-16 and other Mutator proteins in each of the mut-16 deletion backgrounds.
All images are from the transition zone (leptotene/zygotene) region of the germline. Scale bars,
5µm.
75

Figure S5. Regions B and C of MUT-16 are required for interaction with MUT-2 in
vitro.
Immunoprecipitation and western blot of MUT-16::mCherry::2xHA (expected sizes between 132
– 141 kD for MUT-16 deletions and 148 kD for MUT-16 full length) and MUT-2::GFP::3xFLAG (83
kD). Top three panels are total lysate from strains indicated above, and bottom two panels are
following HA immunoprecipitation. Note that in some cases, full-length MUT-16 was degraded
beyond the detection limit of the western blot in the input sample, but was still present following
immunoprecipitation. The equivalent of ~0.5% of starting material for the input fractions and ~20%
of starting material for the IP fractions were loaded onto the gels.

76

Figure S6. MUT-16 foci are temperature dependent.
Representative images from the early pachytene region after L4 MUT-16::GFP animals were
subjected to 30°C heat shock for 6 hours and allowed to recover on plates at room temperature
(~21°C) for the indicated amount of time. All images are from different animals to demonstrate
the variability in foci presence and intensity at each time point. Scale bars, 5µm.

Additional supplemental movies and tables are associated with the original publication.
77
MESO-CHAPTER 1
Phase-separated protein dynamics are affected by fluorescent tag choice.

Associated Publication: Uebel C. J., and C. M. Phillips, 2019 Phase-separated protein
dynamics are affected by fluorescent tag choice. microPublication Biol.
https://doi.org/10.17912/micropub.biology.000143

PREFACE
During my quest to simultaneously visualize the growing number of identified germ
granules, I acquired or created multiple versions of fluorescently tagged P granules. In
my initial experiments with endogenously tagged PGL-1::mKate2, I attempted to dissolve
P granules with 5% 1,6-hexanediol, a treatment which had previously been shown to
disrupt PGL-1::GFP (Updike et al. 2011). Frustratingly, I was unable to reproduce the
results and began to wonder if the fluorescent tag was affecting the liquid properties of P
granules. This work is a result of that initial failed experiment, published in a community-
driven peer-reviewed journal specifically for brief, high-quality data.

ABSTRACT
Techniques to study phase-separated proteins often include fluorescent labeling
and live-imaging microscopy, yet the effects of fluorescent protein tags on the liquid-like
properties of phase-separated condensates is unexplored. In this work, we demonstrate
that the in vivo dynamics of PGL-1 foci are drastically altered when tagged with mKate2
or mTagBFP2 and appear to be more solid or aggregate-like than when tagged with GFP.
Ultimately, our data shows fluorescent tag choice is sufficient to perturb phase separation
dynamics.

78
RESULTS
Biological liquid-liquid phase separation gives rise to dense protein-protein or
protein-RNA condensates that are distinct from the surrounding bulk cytoplasmic or
nuclear phase. These condensates, comprised of many multivalent, weak, and
hydrophobic interactions, perform a wide variety of physiological functions and are
sensitive to changes in the cellular environment (Shin and Brangwynne, 2017). One
notable phase-separated condensate is the P granule, a C. elegans germline-specific
mRNA surveillance center. While the liquid nature of P granules was first described by
Brangwynne et al. (2009), additional P granule properties and protein dynamics have
been examined with a variety of in vivo techniques. The advances in CRISPR/Cas9 gene
editing techniques make it possible to endogenously tag PGL-1, a major constituent of P
granules, and study protein dynamics in vivo via live fluorescent imaging. PGL-1 tagged
with Green Fluorescent Protein (GFP) is widely used for the study of P granules, and
forms distinct perinuclear germline foci consistent with previous observations of P
granules (A, top row) (Pitt et al., 20000; Strome and Wood, 1982).
Due to an interest in visualizing additional germline proteins, we created both pgl-
1::mKate2 (A, middle row) and pgl-1::mTagBFP2 (A, bottom row) endogenously tagged
strains that would allow us to visualize multiple fluorescent proteins with differing
excitation and emission spectra. All three C-terminally fluorescently tagged PGL-1 strains
behave similarly to one another under both undissected and dissected control conditions
(A, B). However, when we tested conditions that probe P granule properties, we were
surprised to find that the mKate2 and mTagBFP2 tagged PGL-1 have strikingly different
dynamics from the previously described PGL-1::GFP.
79
First, 1,6-hexanediol is an aliphatic alcohol that dissolves liquid-like condensates,
but not aggregates or solids (Kroschwald et al., 2017). While the mechanism is not entirely
understood, 1,6-hexanediol is thought to disrupt hydrophobic interactions that are
important for phase-separated condensate integrity. Consistent with Updike et al. (2011),
PGL-1::GFP dissolves in 5% 1,6-hexanediol, as demonstrated by lack of perinuclear foci
and increased fluorescent signal in the cytoplasm (C, top row). In contrast, although PGL-
1::mKate2 and PGL-1::mTagBFP2 have increased cytoplasmic fluorescent signal, clear
perinuclear puncta are still visible, indicating a lack of complete dissolution and perhaps
a more aggregate-like consistency (C, middle row, bottom row).
Second, liquid phase separation is influenced by changes in temperature. Higher
temperatures introduce greater amounts of entropy into the system, allowing for de-
mixing of the condensate into the bulk phase (Alberti et al., 2019). PGL-1::GFP
condensates in embryos are observed to dissolve at temperature shifts to 34°C for 1
minute (Putnam et al., 2019). In adult germlines, evidence of P granule de-mixing can be
seen by 3 hours at 34°C, where PGL-1::GFP foci are faint and fluorescent signal is heavily
cytoplasmic (D, top row). However, this same temperature shift results in large, bright,
abnormal foci and low cytoplasmic signal in both PGL-1::mKate2 and PGL-1::mTagBFP2
(D, middle row, bottom row). The majority of these foci appear detached from the nuclear
periphery and large, round, abnormal aggregates are also observed in the syncytial
gonad rachis (not shown).
Lastly, P granules are protein-RNA condensates, and rely on RNA for their
formation. Consistent with observations by Sheth et al. (2010), PGL-1::GFP foci in the
pachytene are disrupted 5 hours post-microinjection of α-amanitin, a potent
80
transcriptional inhibitor (E, top row). PGL-1::mKate2 and PGL-1::mTagBFP2 both fail to
dissolve, and instead form bright puncta that are abnormal compared to control foci (E,
middle row, bottom row).
Both mKate2 and mTagBFP2 are bright, monomeric fluorescent tags derived from
TagRFP and are similar in size to GFP (Shcherbo et al., 2009; Subach et al., 2011). Here
we show that the in vivo dynamics of PGL-1 foci are drastically altered when tagged with
mKate2 or mTagBFP2 and appear to be more solid or aggregate-like than when tagged
with GFP. While the mechanism behind this effect is unclear, it is possible that specific
tag-to-tag or tag-to-protein interactions are altering the phase dynamics. Additionally, the
high expression levels of PGL-1 may make it more prone to aggregation. Ultimately, our
data shows fluorescent tag choice is sufficient to perturb phase-separation dynamics, and
we recommend ensuring that the dynamics of new fluorescently tagged proteins are
consistent with previous literature, in vitro experiments, untagged protein, or additional
fluorescent tags.

METHODS
Strains
DUP75 pgl-1(sam33[pgl-1::gfp::3xFLAG]) IV
USC1082 pgl-1(cmp145[pgl-1::mKate2::Lox2272::3xMyc]) IV
USC1269 pgl-1(cmp226[pgl-1::mTagBFP2::loxP::3xFLAG]) IV

Strain Construction
The C. elegans wild-type N2 (Bristol) was used as the injection strain for all
constructs. Animals were cultured at 20°C on NGM plates with E. coli (OP50) according
81
to standard condition (S. Brenner, 1974). Fluorescent tags were inserted at the 3’ end of
pgl-1 endogenous locus by CRISPR genome editing using the self-excising cassette as
described (Dickinson et al., 2015). To create the 5’ and 3’ pgl-1 homology arms we used
the following primers:
pgl-1 5’ arm F: GGAGAAGTGTTGTTTGTCCG
pgl-1 5’ arm R: GAAACCTCCACGGCCTCCCCGACCCCCGTAACC
pgl-1 3’ arm F: TAAACTCCAACTATTGAATGTTTAATTTG
pgl-1 3’ arm R: GGCCTCCCTATTAGACTTGC
Silent mutations were included in ‘pgl-1 5’ arm R’ at the site of guide RNA targeting to
protect the plasmid repair template from cleavage. Homology arms also contained 20-
30bp sequences specific to the mTagBFP2 or mKate2 vector for amplification from N2
genomic DNA and subsequent cloning into digested pDD287 (Addgene #70685) for
mKate2 or pJJR81 (Addgene #75029) for mTagBFP2 vectors via isothermal cloning
(Gibson et al., 2009). Correct sequences of constructed plasmids were confirmed with
Sanger sequencing.
The guide RNA for pgl-1 was generated by ligating oligos containing the guide
sequence into BsaI-digested pRB1017 (Addgene #59936) and are the following
sequences for PGL-1: 5’-tcttGGGGGTCGTGGTGGACGCGG-3’ and 5’-
aaacCCGCGTCCACCACGACCCCC-3’. Plasmids were microinjected in the following
concentrations: 50 ng/µL pJW1259 (eft-3::Cas9), 50 ng/µL pgl-1 sgRNA, 25 ng/µL
respective PGL-1 repair plasmid, and 2.5-10 ng/µL GFP or mCherry co-injection markers.
Transgenic animals were confirmed by PCR and are available from the Phillips lab by
request.
82

Microscopy
Adult animals were standardized for age by selection of L4 worms on the day
preceding imaging. Undissected animals were mounted and imaged in <1% sodium azide
in M9 buffer solution to prevent movement. For 1,6-hexanediol experiments, dissected
animals were imaged immediately after dissection in either M9 buffer alone or a solution
of 5% 1,6-hexanediol dissolved in M9 buffer. At least 5 gonads were analyzed for each
genotype. For heat shock experiments, plates of animals were wrapped in Parafilm and
placed in a 34°C incubator for 3 hours. Animals were mounted and imaged in less than
five minutes after removal from incubator. At least 5 animals were analyzed for each
genotype. For transcriptional inhibitor experiments, adult animals were microinjected with
200µg/mL of α-amanitin until the solution flowed around the germline bend. A vehicle
control injection of RNase free water did not disrupt foci. The observed results are
reproducible despite slight variation in microinjected volume and gonad integrity. At least
4 animals were analyzed for each genotype. Due to injection and imaging time
requirements, animals were imaged 5 hours ± 15 minutes post-injection. All live imaging
was performed on a DeltaVision Elite (GE Healthcare) microscope using a 60x N.A. 1.42
oil-immersion objective. Six 0.20 µm Z-stacks were collected and compiled at maximum
intensity projections to create each image. Images were pseudo-colored and
brightness/contrast was increased for clarity using Adobe Photoshop.
83
MESO-CHAPTER 1 FIGURE

Figure 1. PGL-1 fluorescently tagged with mKate2 or mTagBFP2 has distinct
phase-separation dynamics compared to PGL-1::GFP.
(A) Representative live images of endogenously tagged PGL-1::GFP (Green, top row), PGL-
1::mKate2 (red, middle row), and PGL-1::mTagBFP2 (blue, bottom row) expression in the late
pachytene and diplotene region of undissected C. elegans gonads. Scale bar, 10µm. (B) Live
images of endogenously tagged PGL-1 in late pachytene zone of buffer dissected gonad. Scale
bars, 5µm. (C) Live images of late pachytene region of gonads dissected in 5% 1,6-hexanediol,
an aliphatic alcohol. Scale bars, 5 µm. (D) Live images of undissected late pachytene region
immediately after heat shock of 34°C for 3 hours. Scale bars, 5 µm. (E) Live images of pachytene
region 5 hours after microinjection of 200µg/mL of the transcriptional inhibitor, α-amanitin. Scale
bars, 5 µm.

84
CHAPTER 2: MUTATOR FOCI REGULATION
Mutator foci are regulated by developmental stage, RNA, and the cell cycle

Associated Publication: Uebel C. J., D. Agbede, D. C. Wallis, and C. M. Phillips, 2020
Mutator Foci Are Regulated by Developmental Stage, RNA, and the Germline Cell Cycle
in Caenorhabditis elegans. G3 Genes Genomes Genetics 10: g3.401514.2020.
https://doi.org/10.1534/g3.120.401514

PREFACE
I proposed this project during my qualification exams, but did not fully begin to
explore the spatiotemporal regulation of Mutator foci until I started mentoring a high
school student, Dana Agbede, in January 2019. Dana and I initially set out to determine
if germ granules were present in the developmentally quiescent survival-state dauer
larvae of C. elegans, but this endeavor soon became the foundational inspiration for fully
investigating the regulation of Mutator foci.

ABSTRACT
RNA interference is a crucial gene regulatory mechanism in Caenorhabditis
elegans. Phase-separated perinuclear germline compartments called Mutator foci are a
key element of RNAi, ensuring robust gene silencing and transgenerational epigenetic
inheritance. Despite their importance, Mutator foci regulation is not well understood, and
observations of Mutator foci have been largely limited to adult hermaphrodite germlines.
Here we reveal that punctate Mutator foci arise in the progenitor germ cells of early
embryos and persist throughout all larval stages. They are additionally present throughout
the male germline and in the cytoplasm of post-meiotic spermatids, suggestive of a role
in paternal epigenetic inheritance. In the adult germline, transcriptional inhibition results
in a pachytene-specific loss of Mutator foci, indicating that Mutator foci are partially reliant
85
on RNA for their stability. Finally, we demonstrate that Mutator foci intensity is modulated
by the stage of the germline cell cycle and specifically, that Mutator foci are brightest and
most robust in the mitotic cells, transition zone, and late pachytene of adult germlines.
Thus, our data defines several new factors that modulate Mutator foci morphology which
may ultimately have implications for efficacy of RNAi in certain cell stages or
environments.

INTRODUCTION
RNA interference (RNAi) is an evolutionarily conserved strategy to ensure proper
gene expression across a wide range of eukaryotes (Fire et al. 1998; Shabalina and
Koonin 2008). The effectors of RNAi are members of the Argonaute protein family, which
bind small regulatory RNAs ranging from 18-30 nucleotides in length. Together these
components form the RNA Induced Silencing Complex (RISC), which targets fully or
partially complementary transcripts of exogenous or endogenous origin through direct
cleavage, recruitment of exonucleases, or repression of translational complexes
(Hutvagner and Simard 2008; Wu and Belasco 2008). By these means, small RNAs play
key roles in development, fertility, chromosome segregation, and defense against foreign
genetic elements such as transposons and viruses.
In C. elegans, RNA silencing is mediated by a network of ~27 Argonaute proteins
associated with three distinct classes of small RNAs: micro-RNAs (miRNAs), Piwi
interacting-RNAs (piRNAs), and small interfering RNAs (siRNAs). C. elegans siRNAs are
categorized in two distinct groups: primary siRNAs and secondary siRNAs. Primary
siRNAs are cleaved by Dicer from double-stranded RNA and are bound by primary
86
Argonaute proteins like RDE-1 and ERGO-1, whereas secondary siRNAs are produced
from primary siRNA-targeted or piRNA-targeted templates by the RNA-dependent RNA
polymerases (RdRPs), RRF-1 and EGO-1, and are bound by the worm-specific
Argonaute (WAGO) clade of Argonaute proteins (Yigit et al. 2006; Aoki et al. 2007; Pak
and Fire 2007; Sijen et al. 2007; Gu et al. 2009; Vasale et al. 2010). Secondary siRNAs
are crucial for signal amplification and transgenerational silencing; mutant animals that
fail to produce secondary siRNAs display multiple defects including failure to respond to
RNAi, temperature-sensitive sterility, and transposon mobilization (Ketting et al. 1999;
Tabara et al. 1999; Vastenhouw et al. 2003; Zhang et al. 2011).
Secondary siRNA amplification primarily occurs in the mutator complex, which forms
perinuclear foci, known as Mutator foci, in C. elegans germ cells (Phillips et al. 2012).
These Mutator foci are nucleated by MUT-16, which directly interacts with the RdRP RRF-
1, and other key siRNA biogenesis proteins to assemble mutator complexes at these sites
(Phillips et al. 2012; Uebel et al. 2018). The C-terminal region of MUT-16, which contains
regions of intrinsic disorder, is both necessary and sufficient for Mutator foci formation.
Furthermore, Mutator foci behave as phase-separated condensates, with liquid-like
properties such as rapid recovery after photobleaching, temperature and concentration
dependent condensation, and disruption after perturbation of weak hydrophobic
interactions (Uebel et al. 2018). Additionally, Mutator foci are adjacent to P granules,
which are phase-separated mRNA surveillance centers important for maintenance of the
germ cell fate and fertility (Pitt et al. 20000; Brangwynne et al. 2009; Sheth et al. 2010;
Updike et al. 2014; Campbell and Updike 2015; Knutson et al. 2017). Because P granules
also contain proteins associated with the small RNA pathways, including both primary
87
and secondary Argonaute proteins (Wang and Reinke 2008; Claycomb et al. 2009; Gu et
al. 2009; Conine et al. 2010), we hypothesize that P granules and Mutator foci interact at
the nuclear periphery to coordinate small RNA silencing of nascent germline transcripts.
Though P granule regulation and morphology has been extensively studied in all
stages of C. elegans development (Strome and Wood 1982; Updike and Strome 2009),
investigation of Mutator foci is primarily limited to observations within the adult
hermaphrodite germline (Phillips et al. 2012; Uebel et al. 2018). Here, we examine
Mutator foci throughout embryonic, larval, male, and hermaphrodite germline
development and begin to assess factors that regulate or influence Mutator foci
morphology. We determine that Mutator foci first appear as bright, distinct puncta in the
Z2/Z3 progenitor germ cells (PGCs), and that these foci persist throughout all subsequent
developmental stages. We then demonstrate that Mutator foci are present in the male
germline throughout spermatogenesis, and that MUT-16 is sequestered into the
cytoplasm of post-meiotic spermatids. While probing potential regulatory mechanisms for
Mutator foci, we discover that these phase-separated compartments are at least partially
dependent on RNA for their stability in pachytene region of the gonad. Finally, we find
that Mutator foci are largest and most robust in the mitotic cells, the transition zone, and
the late pachytene of adult germlines. By RNAi of key germline development proteins, we
discover that the mitotic cell stage, but not the transition zone, is a determinant of robust
Mutator foci. Thus, through these observations, we better define Mutator foci in all
developmental stages and probe the regulatory factors that influence morphology of this
secondary siRNA amplification center.

88
RESULTS
Robust Mutator foci first appear in the Z2/Z3 progenitor germ cells in C. elegans
embryos
Current characterization of Mutator foci is largely restricted to observations within the
adult germline, without consideration for earlier stages of development. C. elegans
embryos undergo invariant cell divisions, the first of which gives rise to the asymmetric
AB and P1 blastomeres. The P1 cell subsequently produces the P2, P3, and P4 cells, the
latter of which divides around the 100-cell stage to form the Z2 and Z3 PGCs, which
undergo no further divisions until hatching and feeding. After hatching, the PCGs
eventually give rise to the ~2,000 cells that comprise the adult germline. Recent imaging
by Ouyang et al. (2019) observes punctate Mutator foci in the PGCs of later comma-stage
embryos (>550 cells), which describes the onset of elongation around 400 minutes post
fertilization. To determine the earliest formation of Mutator foci, defined here as distinctly
punctate MUT-16 fluorescence, we imaged endogenous MUT-16::mCherry and PGL-
1::GFP in fixed embryos at representative developmental stages. Since MUT-16 is
required for the localization of all known mutator complex proteins, we use MUT-16 foci
as a proxy for Mutator foci, and use the terms interchangeably. At the 2-cell stage, P
granules segregate to the cytoplasm of the P1 blastomere, while MUT-16 predominantly
appears as diffuse cytoplasmic signal in both AB and P1 cells (Figure 1A and A’). At the
16-cell stage, P granules begin to associate with the nuclear periphery of the P4 cell
(Updike and Strome 2010), yet MUT-16 remains diffusely cytoplasmic in all cells of the
embryo (Figure 1B and B’). As the embryos reach the 30-cell stage, coinciding with the
beginning of gastrulation (Sulston et al. 1983), MUT-16 continues to be expressed
throughout the cytoplasm of all embryonic cells (Figure 1C), though we sometimes
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observe faint punctate Mutator foci adjacent to the large, perinuclear P granules (Figure
1C’ and Figure S1). At this stage, any PGL-1 remaining in the somatic blastomeres is
eliminated via autophagy, temporarily creating small PGL-1 foci throughout the embryo
(Hird et al. 1996; Zhang et al. 2009), yet diffuse MUT-16 expression in the cytoplasm is
unchanged. Interestingly, in addition to the cytoplasmic MUT-16 signal in the 100-cell
stage, we consistently observe punctate Mutator foci surrounding the newly formed Z2
and Z3 PGCs (Figure 1D). The distinct foci are adjacent to P granules and at the nuclear
periphery, comparable to Mutator foci localization in adult germ cells. Since we observe
foci in the 300-cell stage, and Ouyang et al. (2019) observes foci in the later comma
stage, Mutator foci appear to persist throughout later stages of embryonic development
(Figure 1E and E’). Thus, our imaging reveals that diffuse cytoplasmic MUT-16 is present
in all embryonic cells at all stages of embryonic development, and punctate Mutator foci
can form as early as the 30-cell stage, but are most consistent and robust in the Z2/Z3
PGCs.

Mutator foci persist in all larval stages of C. elegans
To expand on our characterization of Mutator foci throughout development, we examined
MUT-16 and PGL-1 expression during all larval stages, including the “survival-state”
dauer stage induced by starvation (Cassada and Russell 1975). In fed L1 larva, the PGCs
are easily identified by perinuclear P granules and, despite intestinal autofluorescence,
MUT-16::mCherry is clearly visible in punctate Mutator foci (Figure 2A). Mutator foci are
present in both early and late L2 stages (Figure 2B-C). Additionally, Mutator foci are
visible in the L2d dauer larva, whose germlines are developmentally quiescent (Figure
2D). Finally, Mutator foci are present in the early L3 developmental stage, at which point
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two “arms” of the gonad begin to form and branch from the central somatic gonad
primordium (Figure 2E). Similar to MUT-16 expression in adults, larval germ cells also
show faint cytoplasmic expression and MUT-16 exclusion from the nucleus. Since
Mutator foci have been observed previously in L4 and adult stages (Phillips et al. 2012),
we can conclude that Mutator foci are present across all stages of larval development.

Mutator foci localizes throughout spermatogenesis and MUT-16 is deposited in
the cytoplasm of spermatids
In addition to an emphasis on adult germlines, current studies of Mutator foci rely almost
exclusively on hermaphrodite germlines for characterization. To fully characterize Mutator
foci in male germlines, we live imaged mut-16::mCherry; pgl-1::gfp males. Mutator foci
appear similar to hermaphrodite germlines throughout the mitotic tip, the transition zone,
and the pachytene region (Figure 3A). However, we observe a divergence in the
localization patterns of P granules and Mutator foci in spermatogenesis. Previous
literature reports that PGL-1 and PGL-3 disassemble in late spermatogenesis, though the
P granule component GLH-1 persists until eventual segregation into residual bodies of
budding spermatids (Gruidl et al. 1996; Amiri et al. 2001; Updike and Strome 2010). While
we observe PGL-1 disassembly in late spermatogenesis, we continue to see punctate
Mutator foci around nuclei with no PGL-1 signal (Figure 3A). Mutator foci were previously
observed to localize to the nuclear periphery despite lack of detectable PGL-1 via a glh-
1/glh-4 RNAi knockdown (Phillips et al. 2012), and our findings support the independent
localization of Mutator foci.
To visualize Mutator foci and corresponding nuclei more clearly throughout
spermatogenesis, we dissected and fixed mut-16::gfp male germlines. Using DAPI-
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stained DNA as a guide for morphological identification of chromatin state, we observe
punctate Mutator foci throughout the condensation zone and into the division zone (Figure
3B). Mutator foci are present around nuclei in the karyosome stage, a poorly understood
state of nuclear compaction thought to promote chromosome organization prior to meiotic
division (Shakes et al. 2009). Because chromosomes are highly condensed in the
karyosome stage and the nuclear envelope size remains unchanged, Mutator foci appear
farther away from the DAPI-stained bodies. Additionally, during late diakinesis and
metaphase, the nuclear envelope begins to break down, which may explain why some
punctate Mutator foci in the division zone are no longer associated with DAPI-stained
nuclei (Figure 3B).
Unexpectedly, we discovered MUT-16::GFP signal in the cytoplasm of post-meiotic
spermatids in a unique granular pattern reminiscent of WAGO-1 localization in spermatids
(Conine et al. 2010) (Figure 3C). The presence of MUT-16 in spermatids carries
implications for paternal inheritance of Mutator components and paternal deposition of
WAGO class 22G-RNAs, which have been shown to rescue fertility of piRNA-depleted
RNAi defective hermaphrodites (Phillips et al. 2015).

RNA influences Mutator foci stability and morphology
Many phase-separated condensates are comprised of multivalent interactions between
RNA and proteins, often referred to as ribonucleoprotein (RNP) granules. Concentration,
secondary structure, and RNA length have been shown to regulate size, interaction, and
formation of RNP granules both in vivo and in vitro (Langdon et al. 2018; Garcia-Jove
Navarro et al. 2019). Previously work demonstrated that the presence of RNA is integral
to sustaining P granule integrity in vivo (Sheth et al. 2010). Five hours after gonadal
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microinjection of a potent RNA Polymerase II transcriptional inhibitor, α-amanitin, P
granules disappeared in a pachytene-specific manner. To determine if Mutator foci also
require RNA for their stability, we microinjected 200 µg/mL α-amanitin into the gonads of
adults expressing fluorescently-tagged MUT-16 and PGL-1. We found that both P
granules and Mutator foci began to disperse by three hours, forming fewer foci in both the
mid- and late-pachytene regions (Figure 4A and 4B). In late pachytene, P granules
appeared rounder and more detached from the nuclear pore environment (Figure 4B).
Mutator foci remain adjacent to P granules despite the overall reduction in foci number.
By five and seven hours post α-amanitin injection, P granules and Mutator foci are
nearly completely dissolved in the mid-pachytene region (Figure 4A). In late pachytene,
P granules and Mutator foci also dissipate with increasing severity (Figure 4B).
Interestingly however, we consistently observe faint Mutator foci remaining at seven
hours, despite no detectible PGL-1 foci. This result suggests that Mutator foci may only
partially rely on RNA for stability.
To further explore the effect of mRNA loss on Mutator foci, we placed L4 animals on
RNAi targeting ama-1, the largest subunit of RNA polymerase II, required for mRNA
transcription (Bird and Riddle 1989). Phenotypes from some RNAi can be first observed
after 24 hours of feeding (Kamath et al. 2000). We did not observe any noticeable
perturbation of either P granules or Mutator foci at 24 hours on ama-1 RNAi, but after 30
hours, PGL-1 granules in mid pachytene were faint and fluorescent signal was largely
cytoplasmic (Figure S2). Similarly, MUT-16 foci dissipate after 30 hours on ama-1 RNAi.
Though it is possible that the inhibition of transcription resulted in reduction of PGL-1,
MUT-16, or other major protein constituents of these granules, the dispersed cytoplasmic
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signal of PGL-1 and MUT-16 is more suggestive of foci loss due to lack of mRNA than
due to lack of protein components. These data support our hypothesis that Mutator foci
at least partially rely on RNA for foci integrity.

Mutator foci intensity is dictated by stage of germ cell progression
To examine other elements of Mutator foci regulation, we directed our attention to germ
cell stage and morphology. The C. elegans germline is a syncytium of cells organized in
two branching “arms” extending from a central somatic uterus. The distal tip of each arm
contains actively dividing mitotic cells that progress through meiotic S phase before
transitioning to the leptotene/zygotene stage of meiosis. In this transition zone, nuclei
appear crescent-shaped as chromosomes polarize to find homologs, a trait that is readily
apparent by DAPI staining. The transition zone is followed by early, mid, and late
pachytene, where crossing over of homologous chromosomes occurs. Cells then
progress proximally through the diplotene and diakinesis stages, where chromosomes
condense and the nuclear envelope begins to break down (Figure S3A). By live and
immunofluorescent imaging of endogenously tagged MUT-16, we noticed the early- and
mid-pachytene regions of adult germlines contain dim Mutator foci, whereas the mitotic
tip, transition zone, and late-pachytene regions have large, bright Mutator foci (Figure
5A). We refer to this quality of brightness as “Mutator foci intensity”. To quantify Mutator
foci intensity, we divided each gonad into 10 equal sections and performed a granule
count on images processed with a uniform threshold (Figure S3B). In this manner, dim
foci fell below the threshold and only bright foci were counted, allowing us to determine
the distribution of the brightest foci throughout the germline. To help delineate meiotic
stage, we utilized a central component of the synaptonemal complex, SYP-1, which loads
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during the transition zone and is present on synapsed chromosomes through the
pachytene region (MacQueen et al. 2002). We discovered that the first two regions of the
gonad, which corresponds to the mitotic zone, contains high percentages of above-
threshold Mutator foci (Figure 5B, regions 1-2). However, Mutator foci intensity peaks in
a single region coinciding with the onset of the transition zone (Figure 5B, region 3). This
peak in Mutator foci intensity is followed by a dramatic reduction in intensity in the regions
corresponding to early and mid pachytene (Figure 5B, regions 4-7). Consistent with our
qualitative observations, the foci-depleted region is followed by an increase in foci
intensity in regions corresponding to the late pachytene (Figure 5B, regions 8-10). Our
quantification shows Mutator foci intensity fluctuates in a consistent bright-dim-bright
pattern within adult wild-type germlines.

Mutator foci intensity is associated with mitotic and not transition zone nuclei
Because the number of above-threshold Mutator foci peaks near the transition zone, we
sought to determine if foci intensity was associated with the polarization of chromosomes
that produces the unique crescent-shaped morphology of transition zone nuclei. To first
examine the effects of an extended transition zone, we utilized a mutation in the him-8
gene, required to promote X chromosome pairing and synapsis (Phillips et al. 2005). Cells
that fail to complete synapsis cannot exit the condensed transition state, resulting in
polarized nuclei well into the pachytene region of the germline. Despite having an
extended transition zone region, the brightest Mutator foci in a him-8 mutant gonad
remain restricted to the mitotic region and beginning of the transition zone, with an overall
pattern similar to wild-type animals (Figure S3C). Next, we aimed to determine if Mutator
foci intensity was affected by eliminating the polarized nuclear morphology in the
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transition zone. CHK-2, an ortholog of human checkpoint kinase 2, is required both for
homolog pairing and spatial reorganization of chromosomes at the onset of pairing; chk-
2 mutant germlines do not contain any polarized transition zone nuclei (MacQueen and
Villeneuve 2001). We knocked down chk-2 by RNAi and found that, though chk-2 RNAi
gonads lack polarized nuclei, the intensity pattern of Mutator foci remains similar to wild-
type, with a peak in intensity at the mitotic to meiotic transition (Figure S3D). Our data
indicates that neither extension nor morphological disruption of the transition zone nuclei
perturbs the intensity pattern of Mutator foci.
Because the mitotic region is associated with some of the highest numbers of above-
threshold Mutator foci, we next sought to investigate whether the mitotic cell state
determines Mutator foci intensity. To this end, we knocked down the KH motif-containing
RNA-binding protein GLD-1 and the poly(A) polymerase GLD-2, which are redundantly
required to promote meiotic entry and, when disrupted, produce a tumorous mitotic
germline (Kadyk and Kimble 1998). Because RNAi knockdown was not completely
penetrant, we observe some nuclei entering meiosis, marked by SYP-1 staining (Figure
5C). Despite this variation in penetrance, the gld-2 gld-1 RNAi gonads consistently
produce a proximal mitotic tumor containing very bright Mutator foci. Quantification
reveals more above-threshold foci in the proximal tumor compared to the distal mitotic
zone. Additionally, no peak of fluorescence is found at the onset of SYP-1 loading, in
contrast with the peak near wild-type transition zones (Figure 5D). In gonads with fewer
meiotic nuclei and a larger mitotic tumor, bright Mutator foci are found more uniformly
throughout the germline (Figure S3E). These observations indicate that the mitotic cell
state is a contributing factor of Mutator foci intensity.
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Finally, we sought to manipulate germline morphology and cell state in additional ways
to test if Mutator foci intensity experienced similar perturbations. We knocked down ATX-
2, which functions to promote germ cell proliferation and prevent premature meiotic entry;
RNAi of atx-2 was previously observed to produce small germlines with truncated mitotic
and transition zones (Maine et al. 2004). As expected, immunostained gonads were
smaller than wild-type and had reduced mitotic tip and transition zones (Figure 5E).
Interestingly, Mutator foci appear bright in the mitotic tip but do not form a distinct intensity
peak preceding the transition zone and, in the ensuing pachytene region, foci generally
appear brighter than in the mid pachytene of wild-type animals. Quantification reveals
that above-threshold foci are more evenly distributed throughout the germline (Figures
5F and S3F). Thus, distribution of above-threshold Mutator foci can also be altered by
general perturbations of germline morphology and cell state.

DISCUSSION
Mutator foci are hubs of secondary siRNA biogenesis in the C. elegans germline.
Here we show that Mutator foci first appear in PGCs beginning around the 100-cell stage
of embryogenesis, and that these foci persist in germ cells through all subsequent
developmental stages. Additionally, we find MUT-16 present in post-meiotic spermatids.
We further demonstrate that both the presence of RNA and the germline cell cycle play
key roles in promoting assembly of Mutator foci. Therefore, our work begins to define both
the developmental stages and regulatory factors that shape Mutator foci integrity and
intensity.

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Mutator foci in embryos
Our first observation of robust Mutator foci occurs around the 100-cell stage, after
the P4 cell divides into the PGCs Z2 and Z3. Interestingly, this timing coincides with the
appearance of Z granules, which are germline-specific phase-separated condensates
required for RNAi inheritance. At this time, Z granules de-mix from P granules to form
adjacent foci (Wan et al. 2018). MUT-16, however, does not appear significantly enriched
in P granules prior to Mutator foci formation and is therefore unlikely to be de-mixing from
P granules, but rather forming foci de novo. The birth of the Z2 and Z3 PGCs is also
marked by the disappearance of MEG-3 and MEG-4, which surround the PGL phase of
P granules in early embryogenesis and are crucial for P granule assembly (Wang et al.
2014). Together, these data suggest that there may be a coordinated reorganization of
germ granule components at this time. While the exact mechanism for this reorganization
remains unknown, the timing coincides with a burst of transcription in the PGCs (Seydoux
and Dunn 1997). It is possible that an increase in mut-16 transcript, and thus protein
levels, leads to the emergence of Mutator foci; however, detectable levels of cytoplasmic
MUT-16 protein can be observed in all cells throughout embryonic development.
Therefore, we favor the possibility that newly synthesized transcripts emerging from the
nuclear pores necessitate and perhaps directly promote assembly of the P granule/Z
granule/Mutator foci multi-condensate structures.

Regulation of Mutator foci by RNA
Because the appearance of Mutator foci in embryos coincides with the onset of
transcription in the PGCs, we investigated the effect of transcriptional inhibition in the
adult germline. It was previously shown that injection of a transcriptional inhibitor causes
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pachytene-specific loss of P granules (Sheth et al. 2010). We found that Mutator foci
similarly rely on RNA for their stability in the mid-pachytene region. However, in late
pachytene we observed differences in the dissolution of Mutator foci and P granules
following transcriptional inhibition. Specifically, we observed persistent Mutator foci in the
late pachytene, despite visible dissipation of all P granules at seven hours post-
transcriptional inhibition. The sustained presence of Mutator foci in late pachytene, but
not mid pachytene, could arise for multiple reasons. One possibility is that Mutator foci
may be interacting with more stable classes of RNAs. A recent discovery shows that long
stretches of alternating uridine (U) and guanosine (G), termed polyUG (pUG) tails, are
added to siRNA-targeted transcripts to mark them as templates for secondary siRNA
synthesis (Shukla et al. 2020). These pUG tails, added by the Mutator complex protein
MUT-2, are hypothesized to confer stability to targeted transcripts. If these stabilized
transcripts are associating with Mutator foci in higher quantities in late pachytene, it may
explain the resistance of late-pachytene Mutator foci to dissolution after transcriptional
inhibition. An alternate explanation may reside in the phase-separation properties of
Mutator foci. We previously tested Fluorescence Recovery After Photobleaching on
Mutator foci in the late-pachytene region, and showed that, although there was rapid
recovery of a highly-mobile fraction within Mutator foci, there was also a large non-mobile
fraction of MUT-16 that failed to recover after photobleaching. We have not yet identified
the molecular basis of these distinct phases within Mutator foci, but phase separated
granules can be regulated by post-translational modifications (Li et al. 2013; Wang et al.
2014), making this a tantalizing avenue for future exploration. Nonetheless, this non-
mobile fraction may be in a more gel-like state and therefore resistant to dissipation after
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transcriptional inhibition. A direct comparison of the liquid properties of mid- versus late-
pachytene Mutator foci is necessary to address this possibility.
We also noticed that P granules became more rounded at three hours post-
transcriptional inhibition in late pachytene compared to mid pachytene. Typically, P
granules are intimately associated with nuclear pores, likely through the interaction of the
FG-repeats in DEAD-box helicases GLH-1, GLH-2, and GLH-4, with nuclear pore-
containing FG-repeat proteins (Updike et al. 2011; Marnik et al. 2019). This non-spherical
appearance of P granules due to association with nuclear pores has been described as
a “wetting” of the nuclear membrane (Brangwynne et al. 2009). Interestingly, a mutation
in GLH-1 also creates large, round P granules, possibly caused by an inability to release
captured RNA (Marnik et al. 2019). As first proposed by Sheth et al. (2010), our data
suggests that P granule association with the nuclear periphery may also depend on the
sustained presence of transcripts exiting the nucleus.

Mutator foci in the cell cycle and inheritance
Finally, we explored why Mutator foci brightness varies within different regions of the
germline and determined that perturbations to the cell cycle affected the intensity and
distribution of Mutator foci. We found that nuclei in mitosis tend to be associated with
large, bright Mutator foci, whereas Mutator foci in the early and mid-pachytene are much
dimmer. The exact mechanism governing Mutator foci robustness remains elusive.
Interestingly, the mid-pachytene region is also where Mutator foci are most sensitive to
transcriptional inhibition, again suggesting that more stable RNAs, or perhaps higher
concentrations of small RNA-target transcripts, in the mitotic and late-pachytene regions
may promote condensation of larger foci.
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It has been well-documented that small RNAs can be inherited through both the
maternal and paternal germline (Grishok et al. 20000; Alcazar et al. 2008; Lev et al. 2019).
This transgenerational epigenetic inheritance can promote a memory of self and non-self
transcripts across generations as well as transmit gene regulatory information in response
to environmental conditions (Ashe et al. 2012; Luteijn et al. 2012; Shirayama et al. 2012;
Conine et al. 2013; Rechavi et al. 2014; de Albuquerque et al. 2015; Phillips et al. 2015).
While the Argonaute WAGO-4 and helicase ZNFX-1 have been implicated in the
transmission of maternal small RNAs (Ishidate et al. 2018; Wan et al. 2018; Xu et al.
2018), little is known about how small RNAs are packaged and transmitted through
paternal lineages. Here, we observed Mutator foci throughout spermatogenesis and
detected MUT-16 in the cytoplasm of post-meiotic spermatids. A similarly granular
expression pattern in spermatids was seen for WAGO-1 (Conine et al. 2010); if these
granules coincide, further work will be necessary to determine if they act together to
promote paternal inheritance and whether they promote paternal deposition of not just
small RNAs, but also small-RNA targeted mRNAs. Ultimately, Mutator foci morphology
and regulation may influence efficacy of RNAi in certain cell stages or environments, an
avenue that warrants further investigation.
MATERIALS AND METHODS
Strains
Worms were grown at 20°C according to standard conditions (S. Brenner 1974). Strains
used in this study include:
USC1266 mut-16(cmp41[mut-16::mCherry::2xHA + loxP]) I; pgl-1(sam33[pgl-
1::gfp::3xFLAG]) IV
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USC717 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP]) I (Uebel et al. 2018)
USC1385 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP]) I; him-8(tm611) IV
USC1266 was created by crossing DUP75 (pgl-1(sam33[pgl-1::gfp::3xFLAG]) (Andralojc
et al. 2017) and USC896 (mut-16(cmp41[mut-16::mCherry::2xHA + loxP]) (Uebel et al.
2018; Nguyen and Phillips 2020). USC1385 was created by crossing USC717 (mut-
16(cmp3[mut-16::gfp::3xFLAG + loxP])) (Uebel et al. 2018) and CA257 (him-8(tm611))
(Phillips et al. 2005).

Antibody staining and microscopy
For fixed embryo imaging, gravid adult mut-16::mCherry::2xHA; pgl-1::gfp::3xFLAG
animals were placed in 4 µL water on SuperFrost slides and burst by application of a
glass coverslip to release embryos. Slides were placed on a dry-ice cooled aluminum
block for freeze-crack permeabilization. After coverslip removal, slides were fixed in 100%
ice-cold Methanol for 15 minutes and washed three times in 1xPBST for 5 minutes each.
DAPI was added to the first wash. Embryos were mounted in 10 µL NPG-Glycerol
mounting medium and imaged (Phillips et al. 2009). Embryos were staged by
approximate cell counts and position of the Z2/Z3 PGCs. To minimize artifacts, no
antibody staining was used, as the proteins of interest are fluorescently tagged at their
endogenous loci. To avoid FRET activation and channel bleed-through, 20 Z-stacks were
captured first with 542 nm (red), followed by 475 nm (green) and 390 nm (blue) excitation.
For live imaging, undissected larva, hermaphrodites, or adult males were mounted in
M9 containing <1% sodium azide to inhibit movement or in M9 with no sodium azide, and
images were collected first with 542 nm excitation followed by 475 nm laser excitation.
Larval images were compiled from 10 Z-stacks and staged by gonad morphology. Male
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germline live images were compiled from 20 Z-stacks. Adult hermaphrodite germline live
images were compiled from 5 Z-stacks.
For immunostained germlines, adult males or hermaphrodites were dissected in egg
buffer containing 0.1% Tween-20 and fixed in 1% formaldehyde as described (Phillips et
al. 2009). mut-16::GFP::3xFLAG; him-8(tm611) germlines were immunostained with
1:500 rabbit anti-GFP (Thermo Fisher A-11122), and all other germlines were stained
with 1:2000 mouse anti-FLAG (Sigma F1804) and 1:250 rabbit anti-SYP-1 (Macqueen et
al. 2002). Fluorescent Alexa-Fluor secondary antibodies were used at 1:1000 (Thermo
Fisher). Immunostained germlines were compiled from 25 Z-stacks.
All imaging was performed on a DeltaVision Elite (GE Healthcare) microscope using
a 60x N.A. 1.42 oil-immersion objective. For all images, 0.2 µm Z-stacks were compiled
as maximum intensity projections and pseudo-colored using Adobe Photoshop. Image
brightness and contrast were adjusted for clarity.

Transcriptional Inhibition
Anterior gonads of young adult (~24 hours post L4) hermaphrodites were microinjected
with 200 µg/mL α-amanitin until the solution flowed around the germline bend, as
previously described (Sheth et al. 2010; Uebel and Phillips 2019). A vehicle control
injection of RNase-free water did not disrupt foci, nor were foci disrupted in the uninjected
posterior gonad. The observed results were reproducible despite slight variation in
microinjected volume and gonad integrity. Due to injection and imaging time
requirements, animals were imaged at three, five, and seven hours ±15 minutes post-
injection. At least three animals were imaged for each time point.

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RNA interference
For all RNAi experiments, sequence-confirmed RNAi clones gld-1, gld-2, chk-2, atx-2,
ama-1 and L4440 (control) in HT115 (DE3) bacteria were grown overnight at 37°C to
maximum density. Cultures were concentrated 10-fold and 100 µL was plated on NGM
plates with 5 mM IPTG and 100 µg/mL Ampicillin. For gld-2 gld-1 double RNAi, the
concentrated cell cultures were combined at equal volume before plating. Plates were
stored at room temperature for at least 24 hours to allow for IPTG induction. Synchronized
L1 worms were then plated and grown at 20°C for 70 hours before dissection and imaging,
unless otherwise stated. A minimum of four gonads for each RNAi treatment were
analyzed. RNAi experiments were performed twice to ensure consistent phenotypes.

Quantification of Mutator foci intensity
Gonads were divided by length into 10 equal regions ending at the first single-file
diplotene/diakinesis nucleus. Raw TIFs were loaded into FIJI and backgrounds were
subtracted with a 50-pixel rolling ball radius. Each image was then thresholded identically,
creating a binary image displaying only above-threshold foci. Each region was processed
using the “analyze particles” function and the resulting particle quantification data was
graphed and analyzed in Excel.

ACKNOWLEDGEMENTS
We thank the members of the Phillips lab for helpful discussions and feedback on the
manuscript and the Updike lab for granularity quantification protocols. This work was
supported by the National Institute of Health grants R35 GM119656 (to C.M.P.) and T32
GM118289 (to D.C.W.), and the National Science Foundation Graduate Research
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Fellowship Program Grant No. DGE 1418060 (to C.J.U.). CMP is a Pew Scholar in the
Biomedical Sciences supported by the Pew Charitable Trusts (www.pewtrusts.org) and
C.J.U. is a USC Dornsife-funded Chemistry-Biology Interface trainee. D.A. is a student at
Downtown Magnets High School and performed research under the mentorship of C.J.U.
The him-8(tm611) allele was isolated and provided by Shohei Mitani and the Japanese
National BioResource for C. elegans.
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CHAPTER 2 FIGURES

Figure 1. Distinct Mutator foci appear in the Z2/Z3 progenitor germ cells.
(A-E) Distribution of PGL-1::GFP (green) and MUT-16::mCherry (red) in methanol-fixed embryos
of representative stages. DNA is stained with DAPI (blue) for visualization. Scale bars, 15 µm.
(A’-E’) Enlarged insets of boxed areas from merged embryos in A-E. Small MUT-16 puncta are
visible in the 30-cell stage (yellow arrowhead). Distinct MUT-16 foci are consistently visible in
Z2/Z3 cells of both 100- and 300-cell embryos (white arrowheads). Scale bars, 1 µm.
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Figure 2. Mutator foci are present in L1-L3 larval stages.
(A-E) Live imaging of mut-16::mCherry; pgl-1::gfp expression in representative larval stages.
PGL-1::GFP granules (green) associate with germ cells in the developing gonad in all larval
stages. MUT-16::mCherry foci (red) are also present. Merged panels and enlarged insets (dashed
region) show PGL-1 granules and MUT-16 foci interacting at the nuclear periphery. Merge scale
bars, 5 µm. Inset scale bars, 1 µm.
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Figure 3. Mutator foci are present throughout spermatogenesis and are found in
the cytoplasm of spermatids.
(A) Live images of undissected mut-16::mCherry; pgl-1::gfp male germlines. The distal region
(asterisks) is folded over the pachytene region. Inset (dashed box) is enlarged below to show
MUT-16 foci (red) and PGL-1 granule (green) interactions (white arrows). In late
spermatogenesis, PGL-1::GFP signal becomes undetectable, while punctate MUT-16 foci remain
visible (yellow arrows). (B) MUT-16 foci (green) are present through meiotic division in dissected
and immunostained mut-16::gfp::3xFLAG male germlines. Chromosome morphology is visualized
by DAPI (blue) and distinguishes diplotene nuclei, karyosome formation, the division zone, or
post-meiotic (PM) nuclei. (C) MUT-16 localizes in a granular pattern within the cytoplasm of post-
meiotic spermatids. All scale bars, 5 µm.
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Figure 4. Mutator foci integrity partially relies on RNA.
Germlines of young adult mut-16::mCherry; pgl-1::gfp animals were injected with either a vehicle
control or 200 µg/mL α-amanitin, a transcriptional inhibitor. (A) Live images of the mid-pachytene
region at three, five, or seven hours post injection. MUT-16 foci (red) and PGL-1 granules (green)
are unperturbed in vehicle control injections but dissipate with increasing severity over time. (B)
Live images of the late-pachytene region also reveal an increasing severity of dissipation over
time. At seven hours, some MUT-16 foci remain (white arrows) though PGL-1 granules are
absent. Scale bars, 5 µm.
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Figure 5. Mutator foci intensity varies along the adult germline.
(A) Fixed mut-16::gfp::3xFLAG adult germlines were immunostained for MUT-16 (green), SYP-1
(red) and DNA (blue) to visualize Mutator foci intensity pattern along the germline in relation to
mitotic or meiotic cell state. White dashed line indicates approximate region of the transition zone.
(B) Graph plotting percentage of above-threshold foci along 10 equal sections of mut-
16::gfp::3xFLAG germlines. n = 3. (C) gld-2 gld-1 RNAi produces a tumorous mitotic germline.
MUT-16 foci (green) intensity is bright within the proximal mitotic tumor (asterisks). (D)
Quantification of percent above-threshold foci in gld-2 gld-1 RNAi gonads. n = 4. (E) atx-2 RNAi
germline MUT-16 foci (green) intensity is more evenly distributed than wild-type foci intensity. (F)
Quantification of percent above-threshold foci in atx-2 RNAi gonads. n = 4. In all graphs, foci
counts per region are reported as a percentage of the total foci per gonad to normalize for
differences in gonad size and total foci number. However, quantification of total foci counts shows
similar trends for each condition (see Figure S3F). To determine significance, t-tests were
performed on arcsine transformed percentage values. * indicates a p-value ≤ 0.05, ** indicates a
p-value ≤ 0.01. Distal gonad tips are oriented to the left in all images. In all images, SYP-1 (red)
is used as a marker for meiotic progression and DNA is visualized with DAPI (blue). All scale bars,
15 µm.
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Figure S1. Mutator foci are weakly present in the 30-cell stage.
Live image of a 30-cell embryo (yellow outline) and 100-cell embryo (white outline) within an
undissected mut-16::mCherry; pgl-1::gfp adult. PGL-1::GFP (green) granules associate with the
P4 and Z2/Z3 progenitor germ cells (insets). Small MUT-16::mCherry (red) puncta are visible in
the P4 cell of the 30-cell embryo (yellow arrowheads). MUT-16 foci are clearly visible in the 100-
cell stage (white arrowheads). Image is a maximum intensity projection from five Z-stacks. Top
scale bar, 15 µm. Bottom scale bars, 5 µm.
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Figure S2. Mutator foci dissipate after ama-1 RNAi.
(A) Larval L4 mut-16::mCherry; pgl-1::gfp animals were placed on OP50 or on ama-1 RNAi for 30
hours. MUT-16 foci and PGL-1 granules are present in mid pachytene after 30 hours post-L4 on
OP50, but dissipate after 30 hours post-L4 on ama-1 RNAi. Images are maximum intensity
projection of five Z-stacks. Scale bars, 5 µm.
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Figure S3. Variation in Mutator foci intensity across gonad regions.
(A) Schematic of a wild-type adult hermaphrodite gonad. (B) Gonad divided by length into 10
equal regions ending at the first single-file diplotene/diakinesis nucleus (yellow borders). After
identical thresholding, image shows only above-threshold foci (white) which were quantified using
the analyze particles function in FIJI. (C) mut-16::gfp::3xFLAG; him-8(tm611) mutants display an
extended transition zone (white dashed line) indicated by DAPI-visualized transition nuclei
throughout the gonad (white arrows). MUT-16 foci (green) intensity pattern remains similar to wild-
type. (D) chk-2 RNAi disrupts polarization of DAPI-visualized transition zone nuclei (yellow
dashed line). MUT-16 foci (green) intensity pattern remains similar to the wild-type. (E) Double
gld-2 gld-1 RNAi on mut-16::gfp::3xFLAG animals produces a large proximal mitotic tumor
(asterisks). Numerous bright MUT-16 foci (green) are found throughout the mitotic tumor. Some
SYP-1 (red) staining is present in the more distal region (diamond). DAPI-visualized DNA (blue)
reveals an abundance of small mitotic nuclei in the proximal tumor. Note that the complete distal
tip of the gonad is not shown. (F) Quantification of total foci per gonad region in wild-type (n = 3),
gld-2 gld-1 RNAi (n = 4), and atx-2 RNAi (n = 4) germlines. Each line represents one gonad. All
scale bars, 15 µm.

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MESO-CHAPTER 2
SIMR foci are found in the progenitor germ cells of C. elegans embryos

Associated Publication: Uebel C. J., K. I. Manage, and C. M. Phillips, 2021 SIMR foci are
found in the progenitor germ cells of C. elegans embryos. Micropublication Biology 2021:
10.17912/micropub.biology.000374. https://doi.org/10.17912/micropub.biology.000374

PREFACE
Though my last publication established the formation of Mutator foci in embryos,
and previous literature described P granules and Z granules in embryos, no studies had
yet observed SIMR foci during development. By chance, I saw an early embryo with
distinct SIMR foci in an unrelated slide preparation, prompting me to fully explore SIMR
foci appearance and localization in embryos. The following work is a result of that
serendipitous SIMR sighting.

ABSTRACT
RNA interference is a widely conserved mechanism of gene regulation and
silencing across eukaryotes. In C. elegans, RNA silencing is coordinated through
perinuclear nuage containing at least four granules: P granules, Z granules, Mutator foci,
and SIMR foci. Embryonic localization of these granules is known for all except SIMR
foci. Here we establish that SIMR foci first appear at the nuclear periphery in the P4
germline blastomere and become numerous and bright in the Z2 and Z3 progenitor germ
cells. This timing coincides with the appearance or de-mixing of other germline granules,
providing further evidence for coordinated germ granule reorganization.

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RESULTS
Multiple condensates occupy the perinuclear space of C. elegans germ cells,
where they coordinate RNA surveillance to ensure proper gene expression (Lev and
Rechavi 2020; Sundby et al. 2021). The most well-studied of these condensates are P
granules, phase-separated germ granules required for maintenance of germ cell identity
and fertility (Kawasaki et al. 1998; Updike et al. 2014). P granule morphology and
localization is well documented in C. elegans development (Strome et al. 1982). Adjacent
to P granules are Mutator foci, which are nucleated by MUT-16 and required for the
amplification of small interfering RNAs (siRNAs) to create a robust and heritable silencing
signal (Phillips et al. 2012). During development, faint Mutator foci are occasionally seen
in the P4 germline blastomere of 30-cell embryos, but are most robust and numerous in
the Z2 and Z3 progenitor germ cells (PGCs) of 100-cell embryos (Uebel et al. 2020). A
third germline condensate, Z granules, are situated between P granules and Mutator foci
and facilitate transgenerational epigenetic inheritance of silencing signals. Z granule
components ZNFX-1 and WAGO-4 colocalize with P granules in early embryos, but begin
to de-mix from P granules in the Z2/Z3 PGCs to form separate Z granule condensates
(Wan et al. 2018). Lastly, recently discovered SIMR foci also intimately localize within this
cluster of germline granules. SIMR-1, a key component of SIMR foci, is a Tudor domain
protein that mediates production of secondary siRNAs for piwi-interacting RNA (piRNA)-
targeted mRNAs (Manage et al. 2020). While P granule, Z granule, and Mutator foci
localization through embryonic development has been previously described, the
embryonic appearance of SIMR foci is not known.
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Here we use endogenously tagged SIMR-1::mCherry to investigate the embryonic
onset of SIMR foci. We further visualize embryonic P granules with PGL-1::BFP and
Mutator foci with MUT-16::GFP to compare fluorescence and interaction with SIMR foci.
Live imaging of embryos reveals diffuse cytoplasmic expression of SIMR-1 in all stages
(Figure 1A-E), similar to previous observations of MUT-16 expression (Uebel et al. 2020).
Because SIMR foci are present in the germlines of adult hermaphrodites and localize
adjacent to P granules, we focused our analysis on the germline blastomeres and
progenitor germ cells of embryos (Figure 1A’-E’). In the 2-cell embryo, P granules
segregate to the posterior P1 germline blastomere, yet no punctate SIMR foci are present
(Figure 1A, A’). Similarly, no SIMR foci are found in 8-cell embryos as P granules begin
associating with nuclear pores in the P3 germline blastomere (Figure 1B, B’). The 28-cell
embryo yields the first observable SIMR focus adjacent to perinuclear P granules in the
P4 germline blastomere (Figure 1C, C’). While we consistently observe SIMR foci in 28-
to 50-cell embryos (n = 3), these foci are few and faint. Around the 100-cell stage, the P4
cell gives rise to the Z2 and Z3 PGCs, and it is here that we reliably observe bright and
numerous SIMR foci (Figure1D, D’). These bright foci also persist in the Z2/Z3 of late-
stage embryos of 300 or more cells (Figure1E, E’). Our data reveals the previously
unknown embryonic appearance of SIMR foci.
Consistent with their localization in adult germ cells, embryonic SIMR foci appear
adjacent to both P granules and Mutator foci. Interestingly, the appearance of fewer, faint
SIMR foci in the P4 cell and more numerous, bright SIMR foci in the Z2/Z3 PGCs is similar
to the timing of Mutator foci formation in embryos (Uebel et al. 2020). Both the de-mixing
of Z granules and the appearance of robust Mutator foci and SIMR foci in the PGCs
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correlates with the onset of embryonic germline transcription (Seydoux and Dunn 1997;
Wan et al. 2018; Uebel et al. 2020). Taken together, this observation suggests that the
arrival of newly produced mRNAs in the Z2/Z3 PCGs may necessitate or facilitate the
coordinated reorganization of germ granule components for efficient RNA surveillance.

MATERIALS AND METHODS
USC1401 simr-1(cmp15[simr-1::mCherry::2xHA]) mut-16(cmp3[mut-16::gfp::3xFLAG]) I;
pgl-1(cmp226[pgl-1::bfp::3xFLAG]) IV.

Strain Construction
USC1401 was created by crossing USC1269 (pgl-1(cmp226[pgl-1::bfp::3xFLAG]))
(Uebel and Phillips 2019) and USC774 (simr-1(cmp15[simr-1::mCherry::2xHA]) mut-
16(cmp3[mut-16::gfp::3xFLAG + LoxP]) I; unc-119(ed3) III) (outcrossed) (Manage et al.
2020). All strains are available upon request.

Microscopy
Worms were grown at 20°C according to standard conditions (Brenner 1974). Gravid
adult C. elegans were dissected in 10 µL M9 to expose embryos and mounted on a fresh
2% agarose pad for live imaging. At least 3 embryos were observed for each stage. All
images were acquired with a DeltaVision Elite (GE Healthcare) microscope using a 60x
N.A. 1.42 oil-immersion objective. Ten 0.2-micron Z stacks were compiled as maximum
intensity projections and pseudo-colored using Adobe Photoshop to create each image.
The same exposure, acquisition, and pseudo-coloring settings were used for each image.
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MESO-CHAPTER 2 FIGURE

Figure 1. SIMR foci are numerous and bright in the Z2/Z3 progenitor germ cells.
(A-E) Representative live images of embryos expressing PGL-1::BFP (blue), MUT-16::GFP
(green) and SIMR-1::mCherry (red) to visualize P granules, Mutator foci, and SIMR foci,
respectively. Scale bars, 15 µm. For each stage, at least 3 embryos were observed. (A’-E’) Inset
from boxed outline in A-E merge, highlighting the P lineage and progenitor germ cells. Scale bars,
1 µm. (C’) Triangle arrowhead indicates early SIMR focus. (D’-E’) Notched arrowheads indicate
bright, numerous SIMR foci (red) interacting with Mutator foci (green) and P granules (blue).
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CHAPTER 3: NUAGE INTERACTION AND ORGANIZATION
Mutator foci interact with nuage populations containing all known compartments

Manuscript in progress: Uebel C. J., and C. M. Phillips. C. elegans nuage exhibits an
exterior-to-interior organization and is comprised of discrete populations of P granules,
Z granules, SIMR foci, and Mutator foci.

PREFACE
My final work is an attempt to understand RNA surveillance granules as a complete
unit, focusing on the physical configuration and interaction of granules as well as the
trajectory of biomolecules between granules. The more recent discoveries of two germ
granules, Z granules and SIMR foci, also heightened my interest in understanding how
multiple phase-separated condensates interact at the nuclear periphery without co-
mixing. This project was influenced by a chemical biology T32-funded collaboration with
Dr. Fabien Pinaud’s lab, in which we attempted super-resolution STORM microscopy to
visualize Mutator foci at nanometer scale (see Supplemental Chapter 2). Hindered by
COVID-19 shutdowns and technical limitations, I turned to 3D-Structured Illumination
Microscopy in collaboration with USC’s Core Center for Excellence in Nano Imaging to
gain a high-resolution view of granule organization. The deeper I probed different granule
interactions, the more I began to consider the multiple distinct germ granules as different
compartments to a larger perinuclear nuage, all contributing to the same process of
efficient RNA surveillance. Though this project is ongoing, it has already yielded data with
important implications for RNA routing through the different germ granule compartments.

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ABSTRACT
RNA silencing pathways are complex, conserved, and perform widespread, critical
regulatory roles. In C. elegans germlines, RNA surveillance occurs through a series of
perinuclear germ granule compartments — P granules, Z granules, SIMR foci, and
Mutator foci — multiple of which form via phase separation and exhibit liquid-like
properties. Though the functions of individual proteins within these germ granule
compartments are being explored, the physical interaction, spatial organization, and
coordination of germ granule “nuage” as a whole is less understood. Here we find that
key proteins are sufficient for compartment separation, and that the interaction between
compartments is able to be reestablished after perturbation. Using 3D-Structured
Illumination Microscopy, we discover a previously undescribed toroidal P granule
morphology which encircles all known germ granule compartments. Furthermore, we
quantify the stoichiometric relationships between compartments to reveal discrete
populations of compartment interactions, possibly suggesting functional differences
between nuage configurations. Together, our work creates a more spatially and
compositionally accurate model of C. elegans nuage which informs the conceptualization
of RNA silencing through different germ granule compartments.

INTRODUCTION
Thousands of transcripts undergo RNA surveillance to ensure proper gene
expression in C. elegans. Transcript monitoring and silencing is performed by conserved
regulatory small RNAs comprised of three distinct branches: microRNAs (miRNA), piwi-
interacting RNAs (piRNA), and small interfering RNAs (siRNA). Each small RNA branch
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differs in biogenesis, protein interactors, transcript targets, and primary mechanisms of
action, but the fundamental means of RNA surveillance are conserved: small RNAs,
ranging in size from 18-30 nucleotides, are bound by anciently conserved Argonaute
proteins and together target fully or partially complementary RNAs to direct RNA silencing
through transcriptional or post-transcriptional mechanisms (Ketting and Cochella 2020).
Small RNA-directed silencing is efficient. Exogenously introduced dsRNAs are
routed through the siRNA pathway and can silence transcripts to biochemically
undetectable levels (Fire et al. 1998). A reduction in transcript levels can be detected by
6 hours and silencing outcomes can be phenotypically observed by 24 hours (Kamath et
al. 2000; Yang et al. 2021). Silencing signals directed at germline-expressed genes can
also be inherited through multiple generations by siRNA amplification, maternal or
paternal deposition into progeny, and small RNA-directed accumulation of repressive
chromatin marks on targeted loci (Grishok et al. 2000; Alcazar et al. 2008; Burkhart et al.
2011). RNA silencing also functions to ensure proper gene expression and protect against
deleterious transcripts. In brief, miRNAs regulate genes involved in cell patterning,
development, and lifespan (Ambros and Ruvkun 2018), piRNAs engage most germline
genes and can target transposons, including the Tc3 transposon (Das et al. 2008; Batista
et al. 2008; Lee et al. 2012; Bagijn et al. 2012; Shen et al. 2018), and endogenous siRNAs
target additional transposons, gene duplications, pseudogenes, germline genes, and
repetitive elements (Gu et al. 2009; Fischer et al. 2011). Both piRNAs and some siRNAs,
termed “primary” small RNAs, feed into a downstream amplification step which produces
“secondary” siRNAs. The amplification of primary small RNAs to secondary siRNAs
creates a robust, heritable, efficient silencing signal (Grishok et al. 2000; Aoki et al. 2007;
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Pak and Fire 2007; Vasale et al. 2010). Together, small RNA pathways accomplish
widespread regulatory roles and are required for proper development and fertility.
In the germline, the subcellular organization of these complex and interacting small
RNA pathways occurs within perinuclear germ granules. At least four distinct
compartments, collectively referred to here as nuage, facilitate efficient RNA surveillance,
coordinate small RNA amplification, and ensure small RNA inheritance (Pitt et al. 2000;
Sheth et al. 2001; Phillips et al. 2012; Wan et al. 2018; Manage et al. 2020). P granules,
the first discovered C. elegans nuage compartment, contain both nascent transcripts and
key small RNA pathways components including the Argonautes PRG-1, ALG-3, CSR-1,
and WAGO-1 (Sheth et al. 2010; Batista et al. 2008; Claycomb et al. 2009; Gu et al. 2009;
Conine et al. 2010). P granules also contain other small RNA associated factors including
the Dicer-related helicase DRH-3 and the RNA-dependent RNA polymerase (RdRP)
EGO-1, which act together to produce a subset of secondary siRNAs (Gu et al. 2009;
Claycomb et al. 2009). Also localized to P granules are the piRNA-induced silencing-
defective, PID-1, and the RNase, PARN-1, which are required for the synthesis and
processing of piRNAs (de Albuquerque et al. 2014; Tang et al. 2016; Rodrigues et al.
2019). Finally, most nuclear pores (75%) associate with P granules, indicating the
majority of nascent transcripts associate with P granules upon nuclear export (Pitt et al.
2000). The colocalization of key small RNA components and the direct association with
newly exported transcripts positions P granules as a central compartment for RNA
surveillance.
Adjacent to P granules are Mutator foci, which are considered hubs of small RNA
amplification. Mutator foci are nucleated by the intrinsically disordered protein, MUT-16,
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which recruits key secondary siRNA synthesis proteins, including the RdRP RRF-1 and
all mutator complex proteins (Phillips et al. 2012; Uebel et al. 2018). Recent work has
proposed that the mutator complex protein, MUT-2/RDE-3, a nucleoidyl transferase,
marks transcripts for secondary siRNA synthesis by the addition of long 3’ poly-UG (pUG)
repeats (Shuckla et al. 2020). Loss of Mutator foci causes loss of many secondary siRNAs
and results in an inability to respond to exogenous RNAi (Ketting et al. 1999; Zhang et al.
2011; Phillips et al. 2012). Therefore, Mutator foci are a compartment of nuage in which
siRNA amplification is organized.
A third nuage compartment, Z granules, are often found between P granules and
Mutator foci (Wan et al. 2018). Z granules are named after the first identified component,
a conserved RNA helicase, ZNFX-1, which interacts with a number of small RNA proteins
including the RdRP EGO-1, and the Argonaute proteins CSR-1, WAGO-1, and PRG-1
(Ishidate et al. 2018). Notably, Z granules also contain WAGO-4, a secondary Argonaute
required for transmission of secondary siRNAs to progeny (Wan et al. 2018; Xu et al.
2018). Indeed, znfx-1 mutants respond normally to RNAi, but are unable to pass the
silencing signal on to progeny (Wan et al. 2018). Additionally, ZNFX-1, in coordination
with other factors, appears to balance the production of some secondary siRNAs across
transcripts, preventing accumulation of siRNAs at the 5’ end of transcripts (Ishidate et al.
2018). Z granules, therefore, organize the transgenerational inheritance of small RNAs
and balance the distribution of secondary siRNA synthesis across targeted transcripts.
The most recently discovered nuage compartment involved in siRNA pathways are
SIMR foci, of which only two protein localizations are currently known: SIMR-1, an
extended Tudor domain protein, and RSD-2 (Manage et al. 2020). SIMR-1 was first
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identified in a MUT-16 immunoprecipitation, but found to create functionally-independent
foci directly adjacent to Mutator foci. The name SIMR-1 is derived from siRNA-defective
and mortal germline, as simr-1 mutants lose some mutator-dependent siRNAs and
become sterile over multiple generations at elevated temperature (Manage et al. 2020).
Interestingly, simr-1 mutants desilence a piRNA sensor and lose small RNAs specifically
mapping to piRNA targets (Manage et al. 2020). This mutant phenotype, along with a
conserved role of Tudor domain proteins in both piRNA pathways and in protein-protein
interactions (Pek and Kai 2012), leads to the hypothesis that SIMR foci act in part to direct
piRNA target genes to Mutator foci for downstream siRNA amplification (Manage et al.
2020). Furthermore, the colocalizing factor RSD-2 acts in the exogenous siRNA pathway
to ensure efficient RNAi, suggesting that SIMR foci are a compartment of nuage that more
generally mediates the transition between primary and secondary small RNA pathways
(Han et al. 2008; Sakaguchi et al. 2014; Manage et al. 2020).
All four of these compartments are not bound by lipid membranes. In particular, P
granules, Z granules, and Mutator foci have been shown to form via biomolecular phase
separation, a process by which proteins and RNA de-mix from the surrounding bulk phase
to form concentrated, distinct, and often liquid-like condensates held together by many
weak, multivalent interactions. P granules exhibit controlled dissolution and condensation
in embryos, rapid intramolecular rearrangement, and the ability to drip and fuse off nuclei
when a shearing force is applied (Brangwynne et al. 2009). P granules also dissolve
during heat stress and in 1,6-hexanediol, an aliphatic alcohol that disrupts the weak
hydrophobic interactions that promote phase separation (Updike et al. 2011; Putnam et
al. 2019). Z granules are colocalized with P granules until later in development, when they
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de-mix to form separate, adjacent foci, which also exhibit rapid molecular rearrangement
(Wan et al. 2018). Z granules viscosity is also regulated by the piRNA-induced silencing
defective/Z granule surface protein PID-2/ZSP-1 (Wan et al. 20; Placentino et al. 2021).
Finally, Mutator foci are concentration dependent, dissolve during heat stress and
treatment with 1,6-hexanediol, and are able undergo rapid molecular exchange with the
surrounding bulk phase (Uebel et al. 2018). Thus, at least four distinct perinuclear nuage
compartments, multiple of which are phase separated, are involved in RNA surveillance
(Sundby et al. 2021).
The initial characterization of these compartments has demonstrated individual
contributions to small RNA pathway organization, but the physical interaction and spatial
configuration of all four compartments has only been briefly explored with diffraction-
limited microscopy. It is also unclear how biomolecule exchange of RNAs, proteins, or
small RNAs occurs between compartment boundaries. The current incomplete
understanding of the C. elegans nuage assemblage limits formation of a comprehensive
model to describe how multiple phase-separated compartments organize small RNA
pathways and facilitate RNA silencing
Here we demonstrate that Mutator foci and P granule separation is maintained in
ectopic environments and that granule interaction is dynamic and able to be re-
established after perturbation. We use 3D-Structured Illumination Microscopy (SIM) to
further visualize the phase boundary interaction between Mutator foci, P granules, Z
granules, and SIMR foci. We discover a previously uncharacterized toroidal morphology
of P granules, which we term “P granule pocket”, that interacts with all currently known
compartments of nuage and may promote key small RNA pathway biomolecule
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exchange. Finally, we quantify discreet populations of nuage, which appear to exist in a
hierarchical manner. Ultimately our data begins to construct a more accurate model of
how multiple phase-separated condensates organize the small RNA pathways to elicit
efficient silencing.

RESULTS
Granule separation is maintained in an ectopic environment
In the endogenous germline environment, Mutator foci and P granules exist as
separate, yet adjacent, phase-separated perinuclear condensates (Phillips et al. 2012).
We were interested in determining how these two condensates with liquid-like properties
exist adjacently at the nuclear periphery without co-mixing. We first visualized the
juxtaposition between Mutator foci and P granules in the germline with endogenously
expressed MUT-16::mCherry and PGL-1::GFP, respectively (Figure 1A). To determine if
protein properties alone were sufficient to prevent co-mixing between P granules and
Mutator foci, we overexpressed key proteins for the formation of each compartment in the
C. elegans muscle tissue using the myo-3 promoter. We examined MUT-16, which is
required for Mutator foci formation and had previously been shown to form ectopic foci in
muscle cells when overexpressed, and PGL-1, a major constituent of P granules which
forms ectopic foci when overexpressed in intestines (Updike et al. 2011; Uebel et al.
2018). In previous work, we determined that myo-3-driven overexpression of either
mCherry or GFP alone did not form condensates, and thus overexpressed mCherry-
tagged MUT-16 and GFP-tagged PGL-1 (Uebel et al. 2018). Similar to their interaction at
the germline nuclear periphery, the PGL-1 and MUT-16 condensates maintain their
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separate and adjacent relationship in the ectopic muscle environment (Figure 1B). To
ensure that the interaction was not tag-specific, we also overexpressed PGL-1::mKate2
and MUT-16::GFP in muscle cells and obtained the same result (Figure S1A). Thus, the
separate and adjacent relationship between P granules and Mutator foci does not require
germline cellular environment or association with nuclear pores, but instead relies on the
intermolecular interactions or phase-separation properties of key proteins in each
condensate.
To test if the molecular interactions contributing to phase separation differ
significantly between Mutator foci and P granules, we subjected mut-16::mCherry; pgl-
1::gfp gonads to 1,6-hexanediol, an aliphatic alcohol which disrupts weak, hydrophobic
interactions and dissolves phase-separated condensates (Kroschwald et al. 2017). In a
buffer control dissection, P granules and Mutator foci are clearly visible (Figure 1Ci). Both
P granules and Mutator foci dissolve in 10% hexanediol and 5% hexanediol, as previously
observed (Figure 1Cii, Ciii) (Updike et al. 2011; Uebel et al. 2018). We further dilute the
concentration of hexanediol and observe that P granules are present at both 2.5% and
1.25% hexanediol, yet Mutator foci remain dissolved (Figure 1C iv, v). Mutator foci are
present only in very dilute 0.625% hexanediol (Figure 1Cvi). Repeating the experiment
with GFP-tagged Mutator foci yields similar results (Figure S1B). These data indicate that
Mutator foci are more sensitive to perturbations of hydrophobic interactions via
hexanediol than P granules, and this differential reliance on hydrophobicity for phase
separation may be one factor that prevents co-mixing between adjacent condensates.

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P granule and Mutator foci interaction is dynamic
Many phase-separated condensates, including Mutator foci, dissolve with the
addition of heat (Nott et al. 2015; Uebel et al. 2018). We previously discovered that
Mutator foci dissolved by heat stress quickly reassemble as punctate foci during room
temperature recovery (Uebel et al. 2018). To determine if Mutator foci re-establish
adjacency to P granules after heat-stress dissolution, we heat shocked mut-16::mCherry,
pgl-1::gfp animals for 2 hours at 34 °C. Upon immediate imaging after removal from heat,
we discovered that MUT-16 no longer forms punctate Mutator foci, but instead faintly
colocalizes with P granules in enlarged foci (Figure 1Dii). In previous MUT-16::GFP heat-
stress experiments, we observed a similarly enlarged Mutator focus phenotype, but did
not have P granule markers to compare localization (Uebel et al. 2018). The two-hour
heat shock did not completely dissolve P granules, but many detach from the nuclear
membrane and collect in the syncytial gonad core (Figure S1C). This is possibly an
intermediate phenotype, as previous work demonstrates increased dissolution of PGL-1
after 3 hours at 34 °C heat stress (Jud et al. 2008; Uebel and Phillips 2019). By 30- and
60-minutes room temperature recovery post heat shock, MUT-16 is less visibly
colocalized with P granules and MUT-16 cytoplasmic signal remains high (Figure 1Diii-
iv). By 90- and 120-minutes recovery post heat shock, Mutator foci appear strikingly
similar to the non-heat-shocked control in that they are punctate and adjacent to P
granules (Figure 1Di,1Dv-vi). The ability to quickly re-establish granule interaction after
disruption suggests the adjacent relationship between P granules and Mutator foci does
not need to be established at a specific developmental time point and continually
maintained, but rather that Mutator foci may disassemble and reassemble next to
particular P granules.
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Some P granules exhibit a toroidal morphology
To facilitate RNAi, exchange of biomolecules must occur within C. elegans nuage,
but the physical interface between the different phase-separated compartments is not
well understood. We sought to gain a high-resolution view of the interface between
Mutator foci and P granules using 3D-Structured Illumination Microscopy (3D-SIM), which
has nearly two-fold higher resolution than widefield microscopy (Gustafsson et al. 2008).
In the transition zone, 3D-SIM shows Mutator foci and P granules interacting adjacently
as previously observed in widefield microscopy (Figure 2A). Unexpectedly however,
imaging the mid- and late-pachytene regions revealed a previously uncharacterized P
granule morphology; In these pachytene regions, some P granules appear to surround
Mutator foci in a toroidal “donut-like” morphology, which we further refer to as P granule
pockets (Figure 2B-C, arrow, insets). We also observe a gap between Mutator foci and
the encircling P granule, suggesting the two condensates do not directly interact in P
granule pockets (Figure 2C insets), consistent with previous ideas of compartment
interaction (Wan et al. 2018; Manage et al. 2020). After we characterized P granule
pockets at high resolution, we also began to recognize P granule pockets in widefield
microscopy. P granule pockets are visible in widefield live-imaging of undissected
gonads, though the gap between P granules and Mutator foci is not resolved, likely
obscuring previous characterization of P granule pocket morphology (Figure S2A, red
arrows). Because P granule pockets are present in live samples, the unique morphology
does not appear to be a result of gonad fixation. To test if fluorescent protein tags
influence the formation of P granule pockets, we used anti-HA to immunostain mut-
16::SNAP::HA, which contains a SNAP-tag approximately 10 kD smaller than an mCherry
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tag, and also visualized untagged P granules via anti-PGL-1. Subsequent 3D-SIM
imaging revealed that the antibody-visualized P granules still form P granule pockets,
further validating the authenticity of the P granule structures (Figure S2B, red arrows). Of
note, Pitt et al. (2000) briefly describes arch-shaped P granules, which may be related to
P granule pockets. Altogether, our high-resolution imaging reveals a previously
uncharacterized toroidal P granule morphology, in which some P granules encircle
Mutator foci in the mid- and late-pachytene regions.

Nuage compartments exist in distinct populations
During our microscopy, we observed that not all P granules associate with a
Mutator focus. We therefore hypothesized that nuage compartments interact in distinct
populations and sought to determine the molecular stoichiometry between P granules, Z
granules, Mutator foci, and SIMR foci to define nuage populations. To avoid any artifacts
from non-specific antibody binding, we used native fluorescence from endogenously
tagged mut-16::gfp; rfp::znfx-1; pgl-1::bfp, and mut-16::gfp simr-1::mCherry; pgl-1::bfp at
widefield resolution. Quantification of fluorescent signal revealed that each nucleus (n =
30) in the late pachytene is associated with on average 22.4 (± 3.6) P granules (P), 18.4
(± 2.0) Z granules (Z), 12.4 (± 2.3) SIMR foci (S), and 10.8 (± 2.3) Mutator foci (M) (Figure
3A). Interestingly, P granules are more than twice as abundant as Mutator foci, and Z
granules are 1.7 times more abundant, demonstrating that Mutator foci cannot be
associated with all P granules and Z granules. SIMR foci are more similar in quantity to
Mutator foci, yet still have statistically significant higher counts per nucleus (p = 0.007,
Figure 3A). Our quantifications reveal that P granules are the most abundant nuage
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compartment in the germline, closely followed by Z granules, with Mutator foci as the
scarcest nuage compartment.
While our quantification shows the abundance of each nuage compartment
surrounding a nucleus, it is limited in directly addressing the relationships between the
different compartments. To accurately assess compartment associations, we
simultaneously visualized P granules, Z granules, and Mutator foci and manually
evaluated each compartment for proximity to the other nuage compartments. We found
three main populations of P granules (n = 183) (Figure 3B-D). The first population is
comprised of P granules unassociated with any other visible compartment (P, 22%)
(Figure 3B-C). These solitary P granules are generally smaller than other P granules
(Figure 3D, asterisk and S3A). The second population encompasses P granules
associated only with Z granules (PZ, 35%) (Figure 3B-C). This population ranges more
broadly in size, but is generally comprised of medium and large P granules (Figure 3D,
arrowhead and S3A). The third and most prevalent population, constituting 43% of
assessed P granules, associate with both Z granules and Mutator foci (PZM) (Figure 3B-
C). The majority of these P granules were large, and included P granule pocket
conformations (Figure 3D, arrow and S3A). Of note, both Z granules and Mutator foci are
always adjacent to P granules, implying there are no solitary Z granules or Mutator foci
(Figure 3C). To incorporate all known nuage compartments into our analysis, we next
assessed the proximity of SIMR foci (S) to both P granules and Mutator foci, and found
that the majority of SIMR foci are adjacent to both compartments (PSM, 87%) (Figure 3B,
S3B). In corroboration with our nuage compartment quantification, some SIMR foci were
not associated with Mutator foci (PS, 12%). Only one SIMR focus was not associated with
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any other visible granules (Figure 3B-C), however Manage et al. (2020) finds that 100%
of SIMR foci are adjacent to P granules and all SIMR foci are closely associated with Z
granules (100%). Notably, we find that all Mutator foci are associated with SIMR foci
(Figure 3C, Figure S3B). From these relationship ratios, we extrapolate that P granules
associated with Mutator foci also associate with all other known germ granules and that
PZSM constitutes 43% of all P granule populations. Curiously, nuage compartment
quantification and adjacency rates suggest a hierarchy of nuage nucleation, in which P
granules constitute a base granule and possibly promote the subsequent nucleation of Z
granules, SIMR foci, and Mutator foci.

All nuage compartments are arranged within P granule pockets
Because not all P granules form P granule pockets, we quantified P granule pockets per
nucleus visible at widefield resolution and discovered that nuclei in the late pachytene (n
= 18) have on average 3.8 (±1.2) P granule pockets (Figure S3C). Each P granule pocket
(n = 39) associates with a Mutator focus. Therefore, all nuage compartments are present
in P granule pockets. To assess the physical interaction between P granule pockets and
Z granules or SIMR foci, we used SIM to image immunostained rfp::znfx-1; simr-1::gfp
germlines and P granules. SIMR focus size was variable, but we were often able to detect
a gap between SIMR-1 and P granules, indicating their arrangement within P granule
pockets is similar to Mutator foci (Figure 4A, inset). In contrast, we found that Z granules
consistently occupy the entirety of P granule pocket interiors (Figure 4A, inset). This
suggests that P granules directly interact with Z granules, and that Z granules bridge the
gap between P granule pockets and Mutator foci or SIMR foci, similar to previous
descriptions of PZM interaction (Wan et al. 2018; Manage et al. 2020).
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To investigate the flow of RNA through the distinct P granule pocket nuage
organization, we sought to determine which compartments interact with nuclear pores
and, therefore, with newly exported RNA. Pitt et al. (2000) reports that 75% of nuclear
pores are adjacent to P granules. We therefore immunostained nuclear pore complexes,
Z granules, and Mutator foci and imaged in high resolution with 3D-SIM. Nuclear pore
complexes create similar patterns as P granules, consistent with the previously reported
high degree of interaction. As such, some nuclear pores are arranged in a similar
morphology to P granule pockets, with rafts of pores forming an inner pocket (Figure 4B).
Z granules occupy the entire inner space between rafts of pores and appear to have
minimal overlap with nuclear pores (Figure 4B, inset). Mutator foci are innermost still, and,
like their position within P granule pockets, a distinct gap exists between Mutator foci and
the surrounding nuclear pore complexes (Figure 4B, inset). We conclude that Mutator foci
do not interact with nuclear pores and thus are not directly involved in the capture of newly
exported RNAs. Due to consistent exterior-to-interior organization of P granule pockets,
where P granules appear to be the predominant compartment interacting with newly
exported RNAs, we propose that RNA follows a distinct trajectory through the nuage
compartments.

Exchange of RNAi components occurs within P granule pockets
Because all nuage compartments are arranged within P granule pockets, we were
interested in determining if pocket morphology or internal compartment organization
facilitates exchange of RNA silencing biomolecules. We imaged the key secondary
siRNA-associating Argonaute, WAGO-1, with 3D-SIM to determine sub-compartment
localization (Figure 5). WAGO-1 was previously described to localize to P granules but
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recently found to directly interact with ZNFX-1 of Z granules, despite a lack of fluorescent
colocalization evidence (Ishidate et al. 2018). While we noted the previously described
colocalization with P granules, we found that WAGO-1 also localizes to the interior of P
granule pockets (Figure 5Ai-ii). This extension of localization appears to be specific to P
granule pockets and is not observed in the neighboring non-pocket forming P granules
(Figure 5, asterisks). Because our earlier imaging determined that Z granules reside in
the interior of P granule pockets, our microscopy suggests the interaction between ZNFX-
1 and WAGO-1 may preferentially occur within P granule pockets. Thus, exchange of
RNA surveillance biomolecules between nuage compartments may be promoted within
P granule pockets and begins to suggest that differing nuage populations provide distinct
functionality.

DISCUSSION
Physical Organization of Nuage
Previous to our work, C. elegans nuage had not been characterized using super-
resolved microscopy techniques. Our work adds three key details to the physical
organization of C. elegans nuage. First, nuclear pores do not appear to directly colocalize
with Z granules or Mutator foci. Because previous literature demonstrates a direct
interaction between P granules and nuclear pores (Pitt et al. 2000; Sheth et al. 2010), we
therefore extrapolate that P granules are the first and only compartment to directly capture
newly exported RNA (Figure 6). Second, some P granules form a toroidal morphology
which encompasses all known nuage compartments in a consistent exterior-to-interior
organization. This organization is similar to previous findings (Wan et al. 2018; Manage
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et al. 2020), but distinct in that the P granule surrounds a Z granule, which in turn
encompasses both a SIMR focus and Mutator focus (Figure 6). Third, we provide
preliminary evidence that this organization promotes the exchange of key small RNA
pathway biomolecules. The key secondary Argonaute, WAGO-1, appears to extend
localization beyond P granules to within P granule pockets, a region in which we also
show Z granules localization. Though ZNFX-1 interacts with WAGO-1 in a partially RNA-
dependent manner, colocalization has not been previously observed (Ishidate et al. 2018;
Wan et al. 2018). Thus, our work bridges the gap in these findings and demonstrates
WAGO-1 may localize to other nuage compartments specifically within P granule pockets.
It will be interesting to determine if the exchange of other small RNA components or
targeted transcripts is also promoted within P granule pockets.
The functionality of P granule pockets is an ongoing investigation. New data from
Yang et al. (2021) suggests accumulation of targeted RNA occurs in large, unique P
granules which form between 6 and 21 hours after RNA interference. It will be interesting
to determine if the P granules they observe exhibit the pocket morphology we describe
here. In future studies, it will also be necessary to determine if P granule pocket
morphology actively promotes RNA surveillance, or if it is simply a physical outcome of
granule interaction.

Nuage Compartment Interaction
We find that the separation of P granule and Mutator compartments does not rely
on components of the germline environment, such as association with the nuclear
periphery or germline mRNAs. Rather, separation of the condensate-promoting proteins,
PGL-1 and MUT-16, can be maintained in an exogenous environment, as suggested by
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our ectopic overexpression experiments. Furthermore, we find that PGL-1 and MUT-16
proteins have drastically different responses to perturbation of weak, hydrophobic
interactions, suggesting that compartment separation could also be conferred by a
contrasting composition of the multivalent interactions that promote phase separation,
such as differences in the amino acid content of intrinsically disordered regions. Finally,
we show that Mutator foci are able to re-form and restore their adjacency to P granules
after disruption. This has exciting implications in that Mutator foci could disassemble and
then reassemble, possibly when needed for siRNA production in coordination with a
particular P granule. Although this finding does not include the other nuage
compartments, it begins to illuminate the mechanisms that may maintain and promote
multiple phase-separated structures as distinct compartments within nuage as a whole.

Functionality of Nuage Populations
Our work reveals discrete populations of perinuclear nuage which we suspect play
distinct functional roles in RNA silencing. We suspect that the population of small, solitary
P granules may be in a transitionary state, possibly recently detached from larger P
granules. Pitt et al. (2000) notes that very small P granules are not associated with nuclear
pores. Taken together, we propose that the population of solitary P granules are not
actively monitoring RNA transcripts, which may also inhibit the association of other nuage
compartments. The remaining nuage populations (PZ, PZS, PZSM) appear to form in a
hierarchical manner, in that the association of Z granules with P granules may promote
the nucleation of SIMR foci and then Mutator foci. As Z granules are required for
transgenerational epigenetic inheritance (Wan et al. 2018; Ishidate et al. 2018), we
speculate that PZ populations are dedicated to maintaining the inherited population of
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small RNAs. Finally, since Mutator foci are required to produce a subset of secondary
siRNAs, we speculate that the PZSM populations actively engage new target transcripts
for silencing and small RNA amplification. Together, our work uncovers new details on
the organization and compartmentalization of C. elegans nuage, leading to advances in
the understanding of how RNA silencing is organized through multiple phase-separated
compartments.

MATERIALS AND METHODS
C. elegans strains
Strain # Genotype Source
USC717 mut-16(cmp3[mut-16::gfp::3xFLAG]) I Uebel et al. 2018
USC1242 unc-119 (ed3) III; cmpEx95 [myo-3p::mut-16::gfp; myo3p::pgl-1::mKate] *
USC1266 mut-16(cmp41[mut-16::mCherry::2xHA]) I; pgl-1(sam33[pgl-1::gfp::3xFLAG]) IV Uebel and
Phillips 2020
USC1315 mut-16(cmp249[SNAP::HA::mut-16]) I *
USC1353 mut-16(cmp259 [2xMYC::bfp::mut-16]) simr-1(cmp112[simr-1::GFP::3xFLAG]) I; znfx-
1(gg634[HA::tagRFP::znfx-1]) II
*
USC1376 wago-1(cmp92[gfp::3xFLAG::wago-1]) mut-16(cmp41[mut-16::mCherry::2xHA]) I Nguyen and
Phillips 2021
USC1392 unc-119 (ed3) III; cmpEx97 [myo-3p::mut-16::mCherry; myo-3p::pgl-1::gfp] *
USC1401 simr-1 (cmp15[simr-1::mCherry::HA]) mut-16(cmp3[mut-16::gfp::3xFLAG]) I; pgl-1
(cmp226[pgl-1::bfp::3xFLAG]) IV
Uebel et al. 2021
USC1409 mut-16(cmp3[mut-16::gfp::3xFLAG]) I; znfx-1(gg634[HA::tagRFP::znfx-1]) II; pgl-1
(cmp226[pgl-1::bfp::3xFLAG]) IV
*
USC1410 mut-16(cmp3[mut-16::gfp::3xFLAG]) I; znfx-1(gg634[HA::tagRFP::znfx-1]) II *

* Indicates strains constructed for this chapter
Primers
List of primers used to construct the strains used in this chapter.
Target Sequence
plasmid backbone + myo-3
promoter F
CGCGATGCATTCGAAGATCTGCCCAacctgtactctatcactgcC
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myo-3 Promoter R TTCTAGATGGATCTAGTGGTCG
let-858 3' UTR F GGATGATCGACGCCaACG
let-858 3' UTR R + plasmid
backbone
CACTGATCTTACTTGCACTTATAATACGACTCActagttttccttcctcctct
myo-3 + NotI + mut-16 5' F CAAACCCACGACCACTAGATCCATCTAGAAGCTAGCTGCGGCCGCATGTCCGA
AAGTGATGATGATTATCC
mut-16 internal R TGCATTGGTTGATTGGCTGG
mut-16 internal F GCTACTGACACTTCCAATACACC
let-858 UTR + NotI +mut-16
HA-tag R
AAAATTCAACGACGTtGGCGTCGATCATCCGCGGCCGCTTAAGCGTAGTCTGGA
ACGTCG
myo-3 + NotI + pgl-1 5' F caaacccacgaccactagatccatctagaagctagcGCGGCCGCATGGAAGTTTCAGATCTACG
GG
let-858 UTR + NotI + FLAG-tag
R
AAAATTCAACGACGTtGGCGTCGATCATCCGCGGCCGCTTActtgtcatcgtcatccttgtaa
tcg
HA::SNAP::mut-16 5' homology
arm F
GCAAATTATCGTTTACTTCTCATTTATTGCCGTTCAAACGTTTCAACCCACTCAAA
TACTCGATATTTGCAGAAAatggacaaagactgcgaaatgaagc
HA::SNAP::mut-16 3' homology
arm R
ttcacgatcagtttccgtatacgaggcgggtggtggaccaacaacgatccccaatggatcgatatattgatcggaAat
AtcCaaCtcCGGataatcatcatcactttcggaCATAGCGTAGTCTGGAACGTCGTATGGGTA
GGCATAATCAGGTACATCATAAGGATAacccagcccaggcttgc
mut-16 5' UTR + 2xMYC-tag +
mTagBFP2 F
ATTTGCAGAAAATGGAGCAAAAGCTCATCTCCGAGGAGGACCTCGAGCAGAAG
TTGATCAGCGAGGAAGACTTGTCCGAACTCATCAAGG
5' mut-16 + linker + 3'
mTagBFP2 R
aatggatcgatatattgatcggaAatAtcCaaCtcCGGataatcatcatcactttcggaCATTCCGGCTCC
GTTGAGCTTGTGTCCGAGC
+ 75 nt homology in mut-16
UTR F
GCAAATTATCGTTTACTTCTCATTTATTGCCGTTCAAACGTTTCAACCCACTCAAA
TACTCGATATTTGCAGAAAATGGAGCAAAAGC
+ 75 nt homology in mut-16 R ttcacgatcagtttccgtatacgaggcgggtggtggaccaacaacgatccccaatggatcgatatattgatcgg
guide for N terminal mut-16
insertion
TGATCGGAGATGTCTAATTC
mut-16 5' UTR + SNAP-tag F GCAAATTATCGTTTACTTCTCATTTATTGCCGTTCAAACGTTTCAACCCACTCAAA
TACTCGATATTTGCAGAAAatggacaaagactgcgaaatgaagc
SNAP-tag + 2xHA + mut-16 R ttcacgatcagtttccgtatacgaggcgggtggtggaccaacaacgatccccaatggatcgatatattgatcggaAat
AtcCaaCtcCGGataatcatcatcactttcggaCATAGCGTAGTCTGGAACGTCGTATGGGTA
GGCATAATCAGGTACATCATAAGGATAacccagcccaggcttgc

Strain Construction
To create ectopic expression plasmids, the pCFJ104 myo-3 promoter (Addgene
#19328), full length mut-16 or pgl-1, and either let-858 generic 3’ UTR or endogenous 3’
UTR were amplified via PCR using primers in Table 1. These amplicons were assembled
into a Spe-1 digested pCFJ151 (Addgene #19330) vector with PCR-amplified GFP from
DUP75 (Andralojc et al. 2017), mCherry from USC896 (Uebel et al. 2018), or mKate
amplified from pDD287 (Addgene #70685) using isothermal assembly (Gibson et al.
138
2009). Correct sequences of constructed plasmids were confirmed with Sanger
sequencing. We generated extra-chromosomal arrays as follows: 10 ng/µl myo-3p::mut-
16::gfp, 10 ng/µl myo-3p::pgl-1::mKate2 and 70 ng/µl pBluescript was injected into
HT1593 unc-119(ed3) III and Unc-rescued animals were selected to create USC1242
(cmpEx95). 10 ng/µl myo3p::mut-16::mCherry::2xHA, 10 ng/µl myo3p::pgl-
1::GFP::3xFLAG, and 70 ng/µl pBluescript was injected into HT1593 unc-119(ed3) III and
Unc-rescued animals were selected to create USC1392 (cmpEx97).
For SNAP-tagged MUT-16 (USC1315), a PCR repair template for CRISPR
genome editing was designed with primers (see list) and injected as described in Paix et
al. 2017. Injection mix was prepared as follows: 2.5 µg/µL Cas9 protein (IDT, Cat#
1081059), 100 ng/µL tracrRNA (IDT, Cat# 1072534), 14 ng/µL dpy-10 crRNA (IDT Alt-R
system), 42 ng/µL gene-specific crRNA (IDT) were incubated at 37 °C for 10 minutes.
Following incubation, 10 pmol of SNAP-tag PCR repair template amplified from pSNAP-
tag plasmid (Addgene #101135) and 110 ng/µL of dpy-10 ssODN repair template were
added. The injection mix was centrifuges at max speed, transferred to a fresh tube, and
microinjected into N2 animals. The dpy-10 co-CRISPR created a dominant roller
phenotype for selection of transgenic animals. Transgenic animals were confirmed by
PCR and Sanger sequencing.
For USC1353 a MYG-tag and mTagBFP2 was N-terminally added to mut-16 by
CRISPR as described (Paix et al. 2017). The same guide RNA from USC1315 was used
and similar homology arms were designed, but with MYC and mTagBFP2 as the repair
template. This construct was injected into strain USC1229 (simr-1(cmp112[simr-
1::GFP::3xFLAG]) I; znfx-1(gg634[HA::tagRFP::znfx-1])) (Manage et al. 2020). Because
139
the BFP::MUT-16 signal bleached during SIM image acquisition, we did not use the 405
nm channel for final image construction.
USC1409 and USC1410 were made through various crossing schemes involving
USC1269 (pgl-1(cmp226[pgl-1::bfp::3xFLAG])) (Uebel and Phillips 2019), USC717 (mut-
16(cmp3[mut-16::gfp::3xFLAG])) and YY1446 (znfx-1(gg634[HA::tagRFP::znfx-1])) (Wan
et al. 2018). All strains were maintained at 15 °C, 20 °C, or room temperature (~21 °C)
unless performing heat stress experiments.

Widefield Microscopy Live Imaging
Young adult animals were standardized for age by selection of the L4 larval stage
on the day preceding imaging. For live imaging, undissected animals were mounted on
glass slides in <1% Sodium Azide in M9 buffer solution to prevent movement. For
dissected gonads, animals were quickly dissected in M9 with a #11 Feather Blade scalpel.
Coverslip edges were sealed with clear nail polish to prevent buffer evaporation. All live
imaging was performed on a GE Healthcare DeltaVision Elite microscope using a 60x NA
1.42 oil-immersion objective. Unless otherwise stated, 5 Z-stacks (0.2 µm Z-step) were
compiled at maximum intensity projections to create each image. Adobe Photoshop was
used to pseudo-color and adjust the brightness/contrast of each image for clarity.

Granule Disruption
For 1,6-hexanediol experiments, young adult animals were dissected in either M9
buffer as a control, or a solution of 10%, 5%, 2.5%, 1.25%, or 0.625% 1,6-hexanediol in
M9 buffer. Animals were imaged within 5 minutes after dissection. At least four gonads
were assessed for each dilution.
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For heat-shock experiments, NGM plates of young adult animals were wrapped in
parafilm and placed in an incubator at 34 °C for 2 hours. For room temperature recovery,
plates were allowed to recover on the benchtop at room temperature (~21 °C) for the
specified time. Images of undissected gonads were collected and imaged within ± 10
minutes from each indicated timepoint. At least 3 gonads were assessed for each time
point.

3D-Structured Illumination Microscopy
3D-SIM was performed at the USC Core Center of Excellence in Nano Imaging
using a GE Healthcare DeltaVision OMX V4 microscope with an Olympus 60x NA 1.42
oil-immersion objective. 3 angles and 5 phases were collected for each Z-stack (0.125 Z-
step). Laser wavelengths of 405 nm, 488 nm, 568 nm, and/or 642 nm were used for
excitation. Image processing was performed with softWoRx software for structured
illumination image reconstruction (Weiner filter = 0.005) and channel alignment. Final
images were created by compiling maximum intensity projections of the reconstructions
in softWoRx. Channels were pseudo-colored and brightness/contrast was adjusted for
clarity in Adobe Photoshop.

Immunostaining
For both fixed widefield and 3D-SIM fluorescent imaging, adult animals were
dissected in egg buffer containing 0.1% Tween-20 and fixed for 3 minutes in a 1% v/v
formaldehyde solution. Animals were permeabilized via freeze-cracking and further fixed
in ice-cold 100% methanol for 1 minute. Slide preparation was then performed as
described in Phillips et al. (2009).
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Antibody staining of fixed germlines was performed with 1:500 Rat anti-HA 3F10
(Roche 11867423001), 1:500 Rabbit anti-GFP (Thermo A-11122), 1:50 Mouse IgM anti-
PGL-1 (DSHB Cat#K76), 1:2000 Mouse IgG anti-FLAG M2 (Sigma F-1804), and 1:5000
Mouse IgG anti-Nup107 (Covance mAb414). Fluorescent secondary antibodies used at
1:1000 include Goat anti-rabbit 488 (Thermo A-11008), Goat anti-mouse IgG 488
(Thermo A-11029), and Goat anti-rat 555 (Thermo A-21434). Secondary antibodies used
at 1:500 include Goat anti-mouse IgM 647 (Thermo A-21238) and Goat anti-mouse IgG
647 (Thermo A21236).

Granule and Adjacency Quantification
P granule, Z granule, and Mutator foci quantification was performed on USC1409.
Animals were dissected in 1x PBS and fixed for 20 minutes by adding 1:1 EM-grade
paraformaldehyde for a 4% final solution. Animals were permeabilized via freeze-
cracking, rinsed twice in 1x PBST and stored in 70% ice-cold Ethanol overnight. The
slides were also treated with a slightly modified Molecular Instruments HCR v3.0 RNA-
FISH “generic sample on slide” protocol to label RNA in the far-red channel (See
Supplemental Chapter 3) (Choi et al. 2018). Gonads were mounted in NPG glycerol and
sealed with nail polish. RNA labeling yielded inconclusive data, and thus the far-red
channel was ignored for granule quantification purposes. Because granule and nuclei
integrity were unaffected by HCR-FISH, the images were repurposed for quantification
measurements. The endogenous fluorescence of each granule was utilized for
quantification to avoid any off-target or background artifacts that could arise from
immunostaining. Widefield fluorescent images were taken on the GE Deltavision Elite and
deconvolved with softWoRx software. 10 complete nuclei from the late-pachytene region
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of 3 gonads were selected at random for quantification for a total n = 30. TIFs for individual
channels of each nucleus were converted to 8-bit files, thresholded in FIJI 3D Object
Counter, and manually inspected to ensure appropriate granule coverage through all Z
stacks. SIMR foci quantification was performed on USC1401 in the same manner.
Granule counts were obtained from FIJI 3D Object Counter and plotted via ggPlot.
Adjacency quantification was performed on 3 randomly selected late pachytene
nuclei from each of the 3 previously assessed germlines in both USC1401 for SIMR
populations and USC1409 for P granule populations (n = 9 nuclei per genotype). To aid
in adjacency visualization, fluorescent signal from each channel was converted into a 3D
object using the FIJI 3D-Viewer plugin. Fluorescent thresholding was the same as for
granule quantification, the resampling factor was set to 1, and the display was set to
“surface”. All three channels were layered into one 3D-viewer image for adjacency
assessment and was cross-checked with the original TIF files. For USC1409, P granules
(n = 183), Z granules (n = 142), and Mutator foci (n = 78) were assessed. Raw
quantification data shows 142 Z granules were adjacent to P granules, and 78 Mutator
foci were adjacent to both P granules and Z granules. There were no solitary Z granules
or Mutator foci. Of all P granules assessed, 6 P granules were associated with 2 Z
granules (PZZ, 3%), 3 P granules had 2 Z granules and 1 Mutator foci (PZZM, 2%) and
1 P granule had 2 Z granules and 2 Mutator foci (PZMZM, 1%). These outliers were
grouped as PZ and PZM, respectively. For USC1401, SIMR foci (n = 117) and Mutator
foci (n = 102) were assessed. 116 SIMR foci were adjacent to P granules, and 102
Mutator foci were adjacent to both SIMR foci and P granules. P granules were not
individually counted for USC1401. P granules for both USC1401 and USC1409 were
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manually assessed from the TIF files for P granule pocket morphology and qualitative
size.
ACKNOWLEDGEMENTS
We would like to thank Kevin Keomanee-Dizon at the USC Core Center of Excellence in
Nano Imaging for his expertise and training with 3D-SIM microscopy. C.J.U. is funded by
the National Science Foundation Graduate Research Fellowship Program (DGE
1418060), the USC Research Enhancement Fellowship, and the Chemical Biology
Interface T32-GM118289 NIH/Dornsife grant. The Phillips lab is supported by R35
GM119656, and by the Pew Charitable Trusts (www.pewtrusts.org).

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CHAPTER 3 FIGURES

(Figure legend on following page)
145
Figure 1. Mutator foci and P granule separation is independent of nuclear
association and can to be reestablished after perturbation.
(A) Widefield immunofluorescence of mut-16::mCherry; pgl-1::gfp germlines shows that
endogenous Mutator foci (MUT-16, magenta), and P granules (PGL-1, green), are adjacent yet
distinct compartments. (B) Ectopically expressed MUT-16::mCherry (magenta) and PGL-1::GFP
(green) driven by the myo-3 muscle-specific promoter create condensates that maintain
separation in the muscle environment. (C) Top: Live images of the transition zone of mut-
16::mCherry; pgl-1::gfp gonads dissected in M9 buffer (Ci) or varying concentrations of 1,6-
hexanediol (Cii-vi). Middle: Mutator foci (MUT-16, red) are dispersed in all concentrations of 1,6-
hexanediol except 0.625%. Numbers indicate how many gonads displayed Mutator foci out of
total assessed gonads. Bottom: P granules (PGL-1, green) are disrupted in only 10% and 5% 1,6-
hexanediol, indicating different sensitivities to perturbation of weak hydrophobic interactions.
Numbers indicate how many gonads displayed P granules out of total assessed gonads. (D) mut-
16::mCherry; pgl-1::gfp animals were subjected to heat stress at 34 °C for 2 hours and allowed to
recover at room temperature (~21 °C) for 2 hours. Top: Representative live images of the
pachytene region were collected before heat stress (Di, no h.s.), immediately after heat stress
(Dii, 0 min recovery), and for every 30 minutes during recovery (Diii-Dvi). Middle: Mutator foci
(MUT-16, red) weakly colocalizes with P granules immediately after heat shock. At 30- and 60-
minutes room temperature recovery, Mutator foci loses colocalization with P granules but remains
dispersed in the cytoplasm. By 90- and 120- minutes room temperature recovery, MUT-16
reappears as separate, punctate foci adjacent to P granules, indicating the interaction is able to
be re-established after perturbation. Numbers indicate gonads displaying punctate Mutator foci
out of total assessed gonads. For Dii, colocalization with PGL-1 was seen in 3/3 gonads. Bottom:
P granules do not completely disperse after 2 hours 34 °C heat stress. Numbers indicate how
many gonads displayed P granules out of total assessed gonads. Scale bars, 5 µm.
146

Figure 2. P granules form unique pocket morphologies in the mid and late
pachytene.

3D-Structured Illumination Microscopy images of immunostained mut-16::mCherry; pgl-1::gfp
germlines display P granules (PGL-1, green) and Mutator foci (MUT-16, magenta). (A) P granule
and Mutator focus are adjacent to one another in the transition zone as previously described. (B)
Some P granules appear to form an arc or circular morphology (arrows) around Mutator foci in
the mid pachytene. (C) A toroidal, “donut-like” P granule morphology is readily apparent in the
late pachytene (insets, arrow). Insets highlight the unique morphology, termed “P granule pocket”.
Each P granule pocket appears to surround a Mutator focus, yet a gap is maintained between the
foci. Images for (A) and (B) are comprised of 10 maximum projection Z-stacks (0.125 µm Z-step).
Image (C) is comprised of 55 maximum-projection Z stacks (0.125 µm Z-step). Scale bars, 5 µm.
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Figure 3. Nuage compartments exhibit a hierarchical stoichiometry.
(A) Violin plot of fluorescently tagged germ granules surrounding nuclei in the late pachytene, with
each dot corresponding to one nucleus (n = 30). Asterisks indicate average foci per nucleus. **
indicates p-value ≤ 0.01, **** indicates p-value ≤ 0.0001. Significance was determined with a two-
tailed equal variance Student’s t-test (B) Manual adjacency quantification from either mut-16::gfp;
rfp::znfx-1; pgl-1::bfp to determine P granule populations (left) or simr-1::gfp; rfp::znfx-1; pgl-1::bfp
to determine SIMR foci populations (right). Overlapping pie charts reveal distinct populations of
granule association. (C) Summary of combined granule stoichiometry indicating the percent that
any one compartment (y-axis) is adjacent to a second compartment (x-axis). Of note, Mutator foci
are always associated with all nuage compartments (PZM, n = 183 and PSM, n = 117), suggesting
a hierarchical assembly of compartments. (D) Representative widefield image of a fixed mut-
16::gfp; rfp::znfx-1; pgl-1::bfp late pachytene nucleus displaying the different P granule
populations: P granule only (P, asterisk), P granule associated with Z granule (PZ, arrowhead), P
granule associated with both Z granule and Mutator focus (PZM, arrow). Scale bar, 1 µm.
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Figure 4. P granule pockets exhibit an exterior-to-interior organization.
(A) Structured illumination of immunostained germlines with endogenously tagged SIMR-1
indicating SIMR foci (blue), and ZNFX-1 marking Z granules (magenta). P granules (green) are
visualized with anti-PGL-1 and the inset highlights a P granule pocket. A Z granule occupies the
entire interior of the P granule pocket and a SIMR focus is innermost still (insets). (B) Structured
illumination of immunostained germlines with endogenously tagged ZNFX-1 (magenta) and MUT-
16, labeling Mutator foci (yellow). Nuclear pore complexes (blue) are visualized with anti-Nup 107
(mAb414). A pocket formed by NPCs is highlighted in the inset. A Z granule occupies the interior
of the pocket and a Mutator focus localizes to the center, not directly interacting with nuclear pores
(insets). Scale bars, 1 µm.
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Figure 5. Exchange of biomolecules is facilitated within P granule pockets.
(A) Structured illumination of immunostained mut-16::mCherry wago-1::gfp germlines shows
localization of WAGO-1 (magenta) and Mutator foci (MUT-16, yellow) with P granules (PGL-1,
blue). Insets highlight representative WAGO-1 interaction with P granules and Mutator foci.
WAGO-1 localization extends to the interior of P granule pockets (arrows) but does not extend
past the P granule boundary (dotted line) in non-pocket forming P granules (asterisk).
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Figure 6. Working model of P granule pocket organization of nuage.
Model of a P granule exhibiting toroidal “pocket” morphology at the periphery of a C. elegans
germ cell nucleus (gray). Nuclear pores (dark gray) interact only with the P granule pocket (teal),
enabling P granules to capture nascent RNA (green/black). The P granule pocket encircles a Z
granule (red) which balances secondary siRNA synthesis across transcripts and is required for
siRNA inheritance (Ishidate et al. 2018; Wan et al. 2018). The Z granule further encompasses
both a SIMR focus (purple), which acts as an intermediate between primary and secondary siRNA
pathways, and a Mutator focus, which is required for secondary siRNA synthesis (orange) (Phillips
et al. 2012; Manage et al. 2020). Key siRNA pathway proteins traverse the boundaries of nuage
compartments in P granule pockets (see figure 5), suggesting P granule pocket organization may
be important for RNA silencing.

RNA
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Figure S1. Ectopic interaction and dissolution of granules is consistent across
multiple fluorescent tags.
(A) Ectopically expressed MUT-16::GFP and PGL-1::mKate driven by the myo-3 muscle specific
promoter form distinct condensates that maintain separation in the muscle. This interaction
appears to be independent of the fluorescent tag choice. (B) Live images of gonads dissected in
M9 buffer or a series of 1,6-hexanediol dilutions reveals punctate MUT-16::GFP foci are present
only in buffer and the most dilute 1,6-hexanediol concentration (0.625%), demonstrating the same
pattern of dissolution as MUT-16::mCherry. (C) Live images of mut-16::mCherry; pgl-1::gfp
animals subjected to 2 hours heat shock at 34° C and allowed to recover at room temperature
(~21 °C) for the indicated times. Immediately after heat stress (0 min recovery) and following 30-
and 60-minute recovery, P granules (green) are detached from the nuclear periphery and collect
in the rachis. Fewer detached P granules appear in the rachis of animals after 90- and 120-
minutes of room temperature recovery. Distal gonad region is oriented to the left (asterisk) with
the proximal oocytes below. Scale bars, 5 µm.
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Figure S2. P granule pocket morphology is visible in widefield live imaging and in
untagged P granules.
(A) P granule pockets (red arrows) are detectable in widefield live imaging of the late pachytene
regions of endogenously tagged mut-16::mCherry; pgl-1::gfp animals. The gap between Mutator
foci (magenta) and P granules (green), which was visible in SIM imaging, is not apparent in
widefield imaging (white arrows). Each P granule pocket is associated with a Mutator focus (blue
arrows). (B) SIM imaging of untagged P granules (green) immunostained with anti-PGL-1 also
reveals P granule pockets (red arrows) indicating fluorescent tags are not influencing P granule
morphology. To avoid additional fluorescent tags, Mutator foci (magenta) were visualized via
SNAP::HA::MUT-16 immunostained with anti-HA. Untagged P granule pockets also associate
with Mutator foci (white arrows, blue arrows).
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Figure S3. Further analysis of nuage compartment populations.
(A) P granules (n = 173) were qualitatively assessed as small, medium, or large in size and
subsequently sorted according to associated nuage compartments (P, PZ, or PZM). 71% of P
granules associated with no other compartments were small. P granules associated with Z
granules were more evenly distributed in size. 81% of P granules associated with all other
granules were large. (B) Representative widefield image of a fixed simr-1::gfp; rfp::znfx-1; pgl-
1::bfp late pachytene nucleus showing a P granule associated with both SIMR foci and Mutator
foci (arrow). Scale bar, 1 µm. (C) Box plot showing quantification of P granule pockets per nucleus
(n = 18) in the late pachytene where each dot corresponds to one nucleus.

154
SUPPLEMENTAL CHAPTER 1
Posttranslational modification of MUT-16

RESULTS
Posttranslational modifications are key regulators of phase separation for a
number of phase-separated condensates (Mitrea and Kriwacki 2016). Notably, P granules
disassembly is promoted by the phosphorylation of key granule proteins (Wang et al.
2014). To investigate the regulation of Mutator foci, I aimed to determine if phase
separation of Mutator foci was influenced by posttranslational modification. To this end, I
immunoprecipitated MUT-16 for Liquid Chromatography Mass Spectrometry analysis
(Figure 1A-B). Peptide coverage was limited to 52.3% of the protein, possibly due to the
P/Q/N-rich and repetitive sequence, particularly in the H-I and J regions of MUT-16
(Figure 1C) (Uebel et al. 2018). However, I uncovered two phosphorylated serine
residues at serine 457 (S457) and serine 884 (S884) (Figure 1B). S457 was consistent
with the PHOSIDA database of posttranslational modifications, which also identified
phosphorylation of the preceeding threonine 456 (T456) and of tyrosine 583 (T583), which
fell within an uncovered region of our sample (Figure 1C, pink arrows) (Gnad et al. 2011).
Interestingly, the previously unidentified S884 was positioned within a region of MUT-16
that was both necessary and sufficient for foci formation (Uebel et al. 2018).
To prevent phosphorylation or constitutively mimic phosphorylation for further
analysis of function, I used the CRISPR-Cas9 gene editing system to replace the
identified phosphorylated residues with either valine (V) or alanine (A), which are unable
to be phosphorylated, creating a T456V/S457A (phospho-null) strain and a S884A
(phospho-null) strain in mut-16::gfp animals. I also replaced the phosphorylated amino
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acids with aspartic acid (D) to mimic constitutive phosphorylation, thus creating
T456D/S457D (phospho-mimetic) and S884D (phospho-mimetic) in mut-16::gfp.
To determine if the phospho-null or phospho-mimic mutations affected the function
of MUT-16, I tested whether or not mutants were defective in RNAi, as a key phenotype
of mut-16 mutants is an inability to respond to exogenous RNAi (Ketting et al. 1999). I
tested both pos-1, a germline-specific RNAi, and lin-29, a somatic-specific RNAi. For wild-
type animals, pos-1 RNAi results in dead F1 embryos and lin-29 causes the animal to
rupture from the vulva, in a burst or protruding vulva (Pvul) phenotype. I found that all
phospho-null and -mimetic strains were able to respond appropriately to RNAi, in that
animals fed pos-1 laid dead eggs and the majority of animals fed lin-29 burst from the
vulva. (Figure 1D). Therefore, the function of MUT-16 is unaffected by these particular
phosphorylation mutants and the phospho-null or -mimic lines are not RNAi defective.
To understand if phosphorylation affects the morphology or phase-separation
characteristics of Mutator foci, I live imaged germlines dissected in both control M9 buffer,
or 5% 1,6-hexanediol, to probe the disruption of weak, hydrophobic interactions
(Kroschwald et al. 2017). To my dismay, I was unable to observe any qualitative
phenotype (data not shown). In the buffer control, phospho-mimic or phospho-null foci
appeared as bright and abundant as wild-type MUT-16::GFP and in the 5% 1,6-
hexanediol foci dissolved as expected for wild-type MUT-16::GFP (Data not shown) (for
wild-type MUT-16::GFP response to 1,6-hexanediol: see Chapter 3 Figure S1).
In part, I suspect the lack of apparent functional, morphological, or regulatory
phenotype is due to limited coverage of posttranslationally modified peptides, preventing
identification and subsequent mutation of all modified peptides. However, the regulation
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of phases via posttranslational modification sometimes appears dose-dependent and
able to be fine-tuned depending upon proportion of modified residues (Mitrea and
Kriwacki 2016). Therefore, the first step towards finalizing this investigation would be to
combine the currently known phospho-null and phospho-mimetic mutants via CRISPR,
as mutations in T456/S457 and S884 are currently in separate strains. Second, The Y583
PHOSIDA-identified residue is currently unmutated and should be added to the fully
mutated strain (Gnad et al. 2011). Finally, additional posttranslational modifications could
be predicted with online databases and prediction software and also mutated. Functional
and behavioral phenotypes may be fully tested when a complete suite of residues likely
to be phosphorylated are mutated to a phospho-null or phospho-mimetic state. These
experiments could include the previously described somatic-specific and germline-
specific RNAi to test MUT-16 functionality, a panel of 1,6-hexanediol dilutions to test the
sensitivity of phospho-null and -mimetic MUT-16 to perturbations of hydrophobic
interactions (See Chapter 3, Figure 1), and, ideally, fluorescence recovery after
photobleaching to determine if the liquid-like properties of MUT-16 are affected by
phosphorylation (Uebel et al. 2018).

MATERIALS AND METHODS
Strains
N2 (wild-type)
NL1810 mut-16(pk710) I
USC717 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP]) I
USC1300 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp245[S884A]) I
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USC1301 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp246[S884D]) I
USC1302 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp251 [T456V,S457A]) I
USC1303 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp247 [T456D,S457D]) I
USC1316 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp248 [T456D,S457D]) I
USC1319 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP], cmp252 [T456V,S457A]) I
Strain Construction
Guide RNAs were designed to internal mut-16 PAM sites nearest to the T456/S457 and
S884 residues and DNA repair oligos were designed with silent mutations to protect the
repair oligo from cleavage.

S884D Repair Oligo TCATAATCAGCAAGTTTGAAGTTTCCGTGTGGATCGTATTCTGGA
GGATTaGGaGGgAAtTGATCAGGgtcTCTTCTTCTTTGTGATGAATG
ATCTTCgTGATAAACTGATCTACTGCCATAATGATTGAATTGTGG
ATTACATCCATA
S884A Repair Oligo TCATAATCAGCAAGTTTGAAGTTTCCGTGTGGATCGTATTCTGGA
GGATTaGGaGGgAAtTGATCAGGTGcTCTTCTTCTTTGTGATGAAT
GATCTTCgTGATAAACTGATCTACTGCCATAATGATTGAATTGTG
GATTACATCCATA
T456D/S457D Repair
Oligo
CTTCATCGCACTTTAATCGAAATACTGATCGGTCAACATCTCGTC
CTCCAaGgGCACCCgaTgaTCCAGTCAATCGTGTGATGGAAACAG
ATCCACTAATGGGCCAAGGCACTT
T456V/S457A Repair
Oligo
CTTCATCGCACTTTAATCGAAATACTGATCGGTCAACATCTCGTC
CTCCAaGgGCACCCgtTgCTCCAGTCAATCGTGTGATGGAAACAG
ATCCACTAATGGGCCAAGGCACT
S884D guide RNA GGATTTGGTGGAAACTGATC
T456D/S457D guide
RNA
GGAGAAGTGGGTGCACGTGG

The injection mix was prepared as follows: 0.25 µg/µL Cas9 protein (IDT, Cat#
1081059), 100 ng/µL tracrRNA (IDT, Cat# 1072534), 14 ng/µL dpy-10 crRNA (IDT Alt-R
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system), 42 ng/µL µg gene-specific crRNA (IDT) were incubated at 37 °C for 10 minutes.
Following incubation, 110 ng/µL of oligo DNA repair template and 110 ng/µL of dpy-10
ssODN repair template were added. The injection mix was centrifuged at max speed,
transferred to a fresh tube, and microinjected into USC717 animals. The dpy-10 co-
CRISPR created a dominant roller phenotype for selection of transgenic animals.
Transgenic animals were confirmed by PCR and Sanger sequencing.
Immunoprecipitation
Immunoprecipitation for Mass Spectrometry analysis was carried out on USC717.
Approximately 1.1 million adult worms were collected in IP buffer and frozen down drop-
wise in liquid nitrogen. For an untagged control, 0.5 million N2 woms were collected.
Samples were ground in liquid-nitrogen-cooled mortars with the addition of IP buffer and
the resulting slurry was spun down at 37,000 xg for 30 minutes at 4 °C. M2 anti-FLAG
agarose beads were rotated in IP buffer for 1 hour at 4 °C. N2 and USC717 samples were
then incubated with M2 anti-FLAG beads for 1 hour at 4 °C and subsequently spun down
for pelleting. Beads were washed 5x by rotating with 1 mL IP buffer for 10 minutes. Once
washed, beads were spun down for a final time and resuspended in a 1:1 volume of 2x
Sample Buffer with BME and boiled at 95 °C for 5 minutes. Samples were vortexed and
spun at max speed and the resulting supernatant was transferred to a fresh tube for
analysis. Samples were loaded onto a 4-12% Invitrogen Bolt Bis-Tris Plus gel in 1x MOPS
running buffer and run at 100 V for 40 minutes and an additional 150 V for 35 minutes.
The gel was stained with Coomassie Blue, incubated overnight, and de-stained in distilled
water for at least 4 hours. To determine the correct protein band, 10 µL of input and 3 µL
of IP were run on a Western blot and labeled with mouse anti-FLAG M2 (1:1,000) and
HRP anti-mouse (1:1;000). MUT-16::GFP::3xFLAG was confirmed as a ~150 kD band.
159
The corresponding band was excised from the Coomassie gel and stored in 200 µL
distilled water.

Mass Spectrometry
Liquid Chromatography Mass Spectrometry was performed by the Taplin
Biological Mass Spectrometry Facility at Harvard Medical School. The excised MUT-
16::GFP::3xFLAG band was submitted for detection of protein modifications.

RNAi analysis
For RNAi assays, animals were fed E. coli expressing dsRNA as described
(Kamath et al. 2003a). For pos-1 RNAi, 4 L1 animals were grown to adults on RNAi and
scored 4 days later for hatching of F1 embryos. Number of hatched eggs were reported
as percent hatching with standard deviation from 3 trials. For lin-29 RNAi, 20 L1 animals
were placed on RNAi and scored 3 days later for either a burst or protruding vulva (Pvul)
phenotype. Scores were reported as percent burst or Pvul with standard deviation from 3
trials. Strain response was compared with previous scoring of wild-type or mut-16(pk710)
from Uebel et al. (2018).

160
SUPPLEMENTAL CHAPTER 1 FIGURES

(Figure legend on following page)
161
Figure 1. MUT-16 is posttranslationally modified.
(A) MUT-16::GFP::3xFLAG was immunoprecipitated with anti-FLAG. Wild-type N2 worms were
used as the control (-). 5µL of Color Prestained Protein Standard Broad Range, 20µL input
samples, and 60µL IP samples were loaded onto a 4-12% Invitrogen Bolt Bis-Tris Plus gel in 1x
MOPS running buffer. Coomassie staining revealed two enriched bands at approximately 250 kD
and 150 kD. (B) To determine the correct protein band,10µL of input and 3µL of IP were run on a
Western blot labeled with anti-FLAG (M2) antibodies. Significant enrichment at 150 kDa band was
observed. Molecular weight of MUT-16::GFP::3xFLAG is 149 kD, indicating the enriched bands
in both the Coomassie gel and the Western blot is MUT-16::GFP (A-B, arrows). (C) Schematic of
IP/MS peptide coverage (blue) indicating the identified phosphorylated residues from our data
(magenta) or the Phosida Posttranslational Modification Database (pink) (Gnad et al. 2011). We
identify a previously unknown phosphorylated S884 residue within the foci-formation region of
MUT-16 (JKL) (Uebel et al. 2018). Numbers and letters above the schematic represent the original
partitioning of MUT-16 for analysis in Uebel et al. (2018). (D) Wild-type animals, mut-16 mutant
(pk710) animals, and all alleles of phospho-null or -mimetic animals were assayed for response
to pos-1 germline specific RNAi, which results in dead embryos, or lin-29 somatic specific RNAi,
which causes animals to burst from the vulva. For pos-1, the number of hatched F1 eggs was
scored. For lin-29 RNAi, animals with burst or protruding vulvas (Pvul) were scored. Scores were
displayed as an average percentage with standard deviation from three trials.

162
SUPPLEMENTAL CHAPTER 2
3D-Stochasitc Optical Reconstruction Microscopy (3D-STORM) of Mutator foci

RESULTS
Mutator foci exist on the nanometer scale at or near the diffraction limit of light (200
nm), limiting our ability to visualize the true size of foci, the membraneless boundary of
foci, and the physical interaction with other nuage compartments. To visualize Mutator
foci in high resolution, I initially collaborated with Dr. Fabien Pinaud to employ 3D-
STORM, a microscopy technique which compiles individual signals from stochastically
“blinking” fluorophores to create sub-diffraction-limited images. 3D-STORM can achieve
a resolution of 20-30 nm in the lateral direction, and up to 50-60 nm in the axial direction
(Huang et al. 2009). Furthermore, the point spread function of each blinking signal informs
the Z-position of that signal, enabling a 3D-reconstruction of the object. Using 3D-
STORM, we aimed to (1) define the true size of Mutator foci, (2) determine the distribution
and density of MUT-16 protein within Mutator foci, and (3) visualize the boundary of
membraneless organelles and interactions with neighboring nuage compartments.
We labeled MUT-16::GFP with a photoswitchable AlexaFluor647 nanobooster and
first imaged with Highly Inclined and Laminated Optical (HILO) sheet microscopy to locate
punctate Mutator foci (Figure 1A). We then deactivated the fluorophores to the dark state
with a 647 nm laser and stochastically reactivated the fluorophores using a continuous
488 nm laser. Individual signals of labeled MUT-16 were localized and compiled with
PALMsiever software (Figure 1B). Preliminary imaging without 3D analysis revealed
single-molecule localization of MUT-16, both dispersed within the cytoplasm and
clustered within foci (Figure 1C-D). The MUT-16 signal does not appear excluded within
a nuclear zone, but this is likely an imaging artifact arising from a morphologically
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imperfect gonad and flattened compilation of MUT-16 signals across multiple Z locations
(Figure 1C). Surprisingly, small clusters of MUT-16, comprised of around 10-20 resolved
fluorescent signals, are abundant within the cytoplasmic fraction of MUT-16 (Figure 1D).
It is possibly that these nano-scale foci create small cytoplasmic centers of siRNA
amplification. It would be interesting to determine if cytoplasmic MUT-16 form nano-foci
within the soma, as somatic RNAi is also mutator dependent, but no somatic Mutator foci
have been observed (See discussion in Chapter 1; Uebel et al. 2018). We achieved 20
nm resolution of a single mutator focus which reveals (1) a non-uniform distribution of
MUT-16 within the focus and (2) a poorly-defined focus boundary (Figure 1E). The poorly
defined boundary may reflect the ability of Mutator foci to rapidly exchange MUT-16
proteins with the cytoplasmic fraction of MUT-16 (Uebel et al. 2018).
To create a 3D model of a mutator focus, we located punctate Mutator foci with
HILO microscopy, imaged stochastic AF647 photoswitching with an astigmatic lens, and
assigned Z values using PALMsiever software (Figure 2A-B). From this image we found
that some Mutator foci were close to 100 nm across, far smaller than previously observed
500 nm diffraction-limited foci (Figure 2B) (Phillips et al. 2012; Uebel et al. 2018). As in
Figure 1, MUT-16 distribution both within foci and the cytoplasm appeared non-uniform
(Figure 2B). A 3D volume model of the largest focus (asterisk) was rendered, revealing a
complex Mutator focus morphology, contrary to the previously-assumed round
morphology (Figure 2C).
Although preliminary 3D-STORM data provided surprising insight to the variation
of Mutator foci size and the non-uniform distribution of MUT-16, we had hoped to expand
the project to both capture enough images for statistical analysis of density and
164
distribution, as well as include multiple compartments of nuage in multicolored STORM.
To begin statistical analysis, we would first need to determine the stoichiometry of
nanobooster labeling to MUT-16::GFP molecules, wherein a 1:1 ratio would indicate one
blinking fluorophore represents one MUT-16 molecule. Next, at least 10-15 gonads with
high-quality signal would need to be collected for cluster analysis (Khater et al. 2020).
Unfortunately, only a few high-quality STORM images were collected after protocol
optimization and before COVID-19 shutdowns.
To image multiple compartments with 3D-STORM, we planned on labeling one
protein with the previously described nanobooster activated by 488 nm and imaged at
647 nm, and one protein with benzylguanine (BG)-JaneliaFluor549, a SNAP-tag
conjugable photoswitchable fluorophore, activated by 405 nm and imaged near 549 nm,
allowing for separation between the excitation and emission spectra of our two labeled
substrates (Grimm et al. 2015). Because the nanobooster would label any GFP-tagged
protein, such as PGL-1::GFP, and because we were interested primarily in the interaction
of Mutator with other compartments, I created a MUT-16::SNAP strain for labeling with
BG-JF549. Initial labeling trials of BG-JF549 unfortunately had very high background and
require further optimization (data not shown).
While the completion of this collaboration was unsuccessful, exciting preliminary
data suggests visible Mutator foci range in size from 100 nm to 500 nm, and that MUT-
16 is non-uniformly distributed, both within foci and the cytoplasm. Future work would
require more data collection for statistical analysis and optimization of BG-JF549 labeling.
Ideally, 3D-STORM would reveal the molecular “boundary” between nuage
compartments to create a more accurate model of interaction, and could be expanded to
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incorporate key small RNA molecules, such as WAGO-1, to determine sub-compartment
localization at the single molecule scale.
MATERIALS AND METHODS
Strains
USC717 mut-16(cmp3[mut-16::GFP::3xFLAG + loxP]) (Uebel et al. 2018)
USC1315 mut-16(cmp249[SNAP::HA::mut-16]) I (see chapter 3)
Gonad Preparation for 3D-STORM
Ten worms were dissected in 30 µL EBT (100 µL 10x Egg Buffer, 10 µL 10%
Tween-20, 890 µL H2O) and fixed with 30 µL EM-grade paraformaldehyde (40 µL 10x
Egg Buffer, 200 µL 16% paraformaldehyde, 160 µL H2O) for 20 minutes. Gonads were
further dissected away from the bodies and transferred to 5 µL of EBT on a SuperFrost
slide via mouth pipetting. A #1 coverslip was placed on top and slides were freeze-
cracked on an aluminum block cooled via dry ice. Coverslips were removed and gonads
were permeabilized for 3 minutes in fresh ice-cold 100% methanol. Slides were washed
in the dark for 3 x 10 minutes in 1x PBST, and blocked for 30 minutes in 1% Bovine Serum
Albumin (BSA). Slides were then incubated in the dark for 1 hour with 50 µL of 1:500
gb2AF-647 nanobooster antibody in 1% BSA block. Slides were washed once in 1x PBST
for 10 minutes and twice in 1x PBS for 10 minutes. Slides were then stored overnight in
1x PBS at 4 °C.

Imaging Buffer and Mounting
A reducing imaging buffer was made by dissolving 1 g of Glucose in 10 mL TN
Buffer (50 mM TRIS-HCL, 10 mM NaCl, pH 7.60) for a 100 mg/mL stock solution and kept
on ice. 1 mL of the Glucose-TN solution was added to 0.5 mg of Glucose Oxidase in an
166
eppi tube. A separate 0.1 mg/10 µL stock of Bovine Liver Catalase was made, 4 µL of
which was added to the Glucose + Glucose Oxidase solution. Finally, 9.8 µL of pure BME
was added to the solution, which was then kept at room temperature. 9.5 µL of the
resulting imaging buffer was added directly on top of the isolated gonads mounted on the
slide. Then 0.5 µL of 1:100 TransFluoSpheres Fluorescent Beads (0.036 µm, #T10711)
in filtered TN buffer were added to the middle of the sample. Slides were gently shaken
to distribute beads. A round #1.5 coverslip was placed on top of the sample, and excess
liquid was wicked away. Coverslips were then completely sealed with clear nail polish to
reduce evaporation.

3D-STORM Acquisition and Processing
STORM microscopy was performed on an inverted Nikon Eclipse Ti Microscope
equipped with a 100 x 1.49 NA Nikon objective. Images were collected by an iXon
EMCCD Andor camera with the astigmatic lens engaged for 3D-reconstruction.
Fluorophore photoswitching was achieved by continuous low power excitation with 488
nm and imaged with the 647 nm laser excitation. Z-calibration for point spread functions
was performed by collecting complete Z-stacks of the TranFluoSpheres and applying
Gaussian fitting using rapidSTORM software as described in Fernandez et al. (2021).
Fluorescent signals were compiled by the PALMsiever program and 3D volume rendered
via a PALMsiever plug-in.

167
ACKNOWLEDGEMENTS
I would like to thank Dr. Fabien Pinaud and Tony Fernandez for this wonderful
collaboration opportunity. Thank you also to Luke Lavis for the generous gift of BG-JF549
and to Aaron Balana of the Pratt Lab for purifying our attempt at conjugated BG-JF549.
This work was funded by the NIH/Dornsife T32-GM118289 Chemical Biology Interface
grant.

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SUPPLEMENTAL CHAPTER 2 FIGURES

Figure 1. Distribution of MUT-16 protein is non-uniform.
(A) Highly Inclined and Laminated Optical sheet (HILO) microscopy of mut-16::gfp in a
fixed gonad labeled with photoswitchable Alexa-Fluor647. (B) Superresolved PALMsiever
compilation of individual signals above intensity threshold in rapid-STORM analysis on
gonad in A. (C) 10-fold zoom of boxed region in B (blue) highlighting a single nucleus. (D)
10-fold zoom of boxed region in C (red) highlighting a single Mutator focus. (E) Zoom of
boxed region in D (yellow).
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Figure 2. Mutator foci have complex morphologies.
(A) Highly Inclined and Laminated Optical sheet (HILO) microscopy of mut-16::gfp in the
transition zone of a fixed gonad labeled with Alexa-Fluor647 photoswitchable
nanobooster. (B) PALMsiever compilation of individual signals above intensity threshold
in rapid-STORM analysis. MUT-16 distribution within foci is non-uniform and focus size
ranges from ~100-500nm. Color bar indicates Z-axis position of individual signals
calculated via point spread function. Scale bar, 100nm. (C) 3D-Volume render of a single
focus (B, asterisk) generated by PALMsiever shows complex morphology not revealed in
widefield imaging. Scale bar, 100nm.
170
SUPPLEMENTAL CHAPTER 3
Visualization of RNA within C. elegans germlines

RESULTS
To create a comprehensive model of RNA silencing through multiple
compartments of nuage, I sought to visualize the localization of RNA targeted by both the
endogenous and exogenous siRNA pathways. To visualize RNA, I adapted the
Hybridization Chain Reaction RNA Fluorescent in situ Hybridization (HCR-RNA-FISH)
protocol and optimized it for gonads dissected on a slide (Choi et al. 2018). To avoid
needing to label protein via immunofluorescent staining, a protocol which uses contrasting
methods to HCR-RNA-FISH, I utilized the native fluorescence of endogenously CRISPR-
tagged simr-1::gfp; rfp::znfx-1; pgl-1::bfp. I first labeled the RNA of Tc1, a transposable
element, and bath-45, a pseudo gene, both of which are heavily silenced by the mutator-
dependent 22G-RNA endogenous pathways (Zhang et al. 2011; Phillips et al. 2012;
Phillips et al 2014). As expected of constitutive targets of the RNA silencing pathway,
levels of both bath-45 and Tc1 RNA are largely undetectable in the cytoplasm (Figure 1A-
B). There is also no obvious localization within germ granules. Interestingly, there appear
to be singlets or doublets of bright RNA foci near or within the nucleus (Figure 1A-B,
insets). Doublet signals are more apparent in the late pachytene, particularly within the
labeled Tc1 RNA images. I suspect these foci mark nascent transcription of bath-45 or
Tc1 loci on homologous chromosomes, but additional microscopy with DAPI-labeled DNA
or DNA-FISH is necessary to confirm this hypothesis. If these foci indeed represent
transcription at the endogenous locus, it is interesting that only one locus appears to be
actively transcribed for the Tc1 element, which is known to exist in at least 30 discrete
locations (Fischer et al. 2003). I speculate that this may be a way to preserve small RNA
171
production for silencing the Tc1 elements, but how a particular locus, out of a multitude
of loci, is marked for sustained transcription is unknown. Yang et al. (2021), suggests that
genetic loci targeted for silencing by exogenous RNAi localize to the inner nuclear
membrane adjacent to large P granules. It would be interesting to determine if the active
Tc1 locus also localizes near large P granules, or perhaps P granule pockets, which
contain all known nuage compartments (see chapter 3). Since nuclei in the late pachytene
contain an average of 3.8 P granule pockets (see chapter 3), it is reasonable to speculate
that constitutively silenced genes could segregate to specialized configurations of nuage
for sustained production of small RNAs from limited RNA transcription.
To visualize the trajectory of exogenously silenced transcripts through nuage, I
placed mut-16::gfp; rfp::znfx-1; pgl-1::bfp animals on oma-1 RNAi to silence germline
specific oma-1 RNA. This experiment was also an attempt to recapitulate the findings by
Yang et al. (2021), to determine if the large P granules they observed were the P granule
pockets we describe in Chapter 3. Yang et al. (2021) suggests that targeted transcripts
begin accumulating in large P granules at 6 hours, and that by 21 hours 100% of nuclei
contain one large P granule which accumulates the RNAi-targeted RNA. Therefore, I
subjected animals to RNAi for 21 hours and used HCR-RNA-FISH to visualize the
localization of oma-1 RNA. Compared to the control, oma-1 levels are considerably
reduced after 21 hours on RNAi (Figure 2A-B). However, I did not observe any obvious
and consistent colocalization with P granules, large or otherwise, or even within any of
the nuage compartments. Because this was contrary to previously published studies
(Yang et al. 2021), I speculate that the method of HCR-RNA-FISH might be preferentially
amplifying easily accessible cytoplasmic RNA rather than the granule-accumulated RNA.
172
To overcome this technical challenge, we aim to employ single molecule RNA FISH in
future studies and have currently started optimizing the protocol. Single molecule RNA-
FISH would also need to be applied for visualization of Tc1 and bath-45 to determine if
the RNA localization we observe with HCR-FISH is accurate. This ongoing work,
particularly if paired with 3D-Structured Illumination Microscopy, would allow us to
visualize the trajectory of RNA through compartments of nuage at various timepoints after
exogenous RNAi, thus enabling us to create a detailed spatiotemporal model of RNA
silencing.

MATERIALS AND METHODS
Strains
USC1296 simr-1(cmp112[simr-1::gfp::3xFLAG + loxP]) I; znfx-
1(gg634[HA::tagRFP::znfx-1]) II; pgl-1(cmp226[pgl-1::BFP +loxP + 3xFLAG]) IV
USC1409 mut-16(cmp3[mut-16::gfp::3xFLAG + loxP]) I; znfx-
1(gg634[HA::tagRFP::znfx-1]) II; pgl-1 (cmp226[pgl-1::bfp::3xFLAG]) IV

Strain construction
USC1296 was created by crossing USC1269 (pgl-1(cmp226[pgl-1::bfp::3xFLAG]))
(Uebel and Phillips 2019], USC1229 (simr-1(cmp112[simr-1::gfp::3xFLAG + loxP]) I; znfx-
1(gg634[HA::tagRFP::znfx-1]) II) (Manage et al. 2020]. Construction of USC1409 is
described in Chapter 3.
HCR-RNA-FISH
RNA probe sets, probe hybridization buffer, probe wash buffer, amplification buffer
and amplifier hairpins were obtained from Molecular Instruments. All dissection, probe
173
hybridization, and buffer exchanges were done with RNase-free reagents. For RNAi
experiments, L4 stage animals were plated on either L4440 control or oma-1 RNAi
prepared as described (Kamath et al. 2003a) and harvested 21 hours later. Animals were
washed in 1x PBS and transferred to 30 µL 1x PBS for dissection with a clean #11
Featherblade scalpel. EM-grade paraformaldehyde was added for a final concentration
of 4% v/v. Dissected gonads were fixed for 20-30 minutes. Excess liquid was aspirated
off and gonads were mounted on SuperFrost slides. Slides were freeze-cracked on an
aluminum block cooled by dry ice and rinsed twice in 1X PBST. Slides were permeabilized
in 100% ice-cold ethanol overnight or for up to 1 week. After permeabilization, 100 µL
hybridization buffer was added directly to the sample for 10 minutes at 37 °C in a
humidified chamber. Slides were incubated overnight at 37 °C in 50 µL of 0.4 pmol of the
appropriate probe set in probe hybridization buffer. Slides were then washed at 37 °C in
250-500 µL of the following: 100% probe wash (rinse), 75% probe wash/25% 5xSSCT
(15 minutes), 50% probe wash/50% 5xSSCT (15 minutes), 25% probe wash/75%
5xSSCT (15 minutes), 100% 5xSSCT (15 minutes). A final wash consisted of room
temperature 100% SSCT for 5 minutes. 4 µL of each 3µM far-red amplification probe H1
and far-red amplification probe H2 were separately heated at 95 °C for 90 seconds and
allowed to snap-cool at room temperature in the dark for 30 minutes. During these 30
minutes, 100 µL of amplification buffer was added directly to the sample for pre-
amplification. Snap-cooled H1 and H2 were then mixed in 200µL amplification buffer and
50 µL of this solution was added to each slide for 45 minutes. Amplification probes were
washed off with a series of 500 µL 5x SSCT washes. Gonads were mounted with 10 µL
NPG-Glycerol and a #1.5 coverslip sealed with clear nail polish.
174
SUPPLEMENTAL CHAPTER 3 FIGURES

Figure 1. Visualization of endogenously silenced RNA.
(A) HCR-RNA-FISH of bath-45 RNA in a fixed simr-1::gfp; rfp::znfx-1; pgl-1::bfp germline. RNA
(red), Z granules (ZNFX-1, yellow), SIMR foci (SIMR-1, blue) and P granules (PGL-1, purple) are
shown in the mid–late pachytene with the distal tip oriented to the left. Insets are from the boxed
region in A (yellow dotted line). (B) HCR-RNA-FISH of Tc1 RNA in a fixed simr-1::gfp; rfp::znfx-1;
pgl-1::bfp germline. RNA (red), Z granules (ZNFX-1, yellow), SIMR foci (SIMR-1, blue) and P
granules (PGL-1, purple) are shown in the mid–late pachytene with the distal tip oriented to the
left. Insets are from the boxed region in B (yellow dotted line). Scale bars, 5 µm.
175

Figure 2. Visualization of oma-1 targeted by RNAi.
(A) HCR-RNA-FISH of oma-1 RNA in fixed mut-16::gfp; rfp::znfx-1; pgl-1::bfp germlines of
animals grown on OP50, placed on L4440 control RNAi at the L4 larval stage, and harvested 21
hours later. RNA (red), Z granules (ZNFX-1, yellow), Mutator foci (MUT-16, blue) and P granules
(PGL-1, purple) are shown in the mid–late pachytene with the distal tip oriented to the left. Insets
are from the boxed region in A (yellow dotted line). (B) HCR-RNA-FISH of oma-1 RNA in fixed
mut-16::gfp; rfp::znfx-1; pgl-1::bfp germlines of animals grown on OP50, placed on oma-1 RNAi
at the L4 larval stage, and harvested 21 hours later. RNA (red), Z granules (ZNFX-1, yellow),
Mutator foci (MUT-16, blue) and P granules (PGL-1, purple) are shown in the mid–late pachytene
with the distal tip oriented to the left. Insets are from the boxed region in A (yellow dotted line).
Images are max intensity projections of 20 Z-stacks (0.2 µm Z-step). Scale bars, 5 µm.
176
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Abstract (if available)

Abstract Small RNA pathways are critical regulators of gene expression in eukaryotic cells. In the nematode C. elegans, small RNAs target and downregulate complementary RNA to ensure proper gene expression and silence deleterious transcripts. Robust silencing is achieved by small RNA amplification, which is dependent upon a perinuclear membraneless compartment in germ cells called Mutator foci. Disruption of Mutator foci causes chromosomal nondisjunction, temperature-sensitive sterility, and transposon activation. ❧ My doctoral work initially investigates the molecular formation of Mutator foci. First, in examining different regions of MUT-16, a protein which nucleates Mutator foci, I discover important functional subdivisions of the MUT-16 protein which are necessary for protein-protein interaction and foci formation. I further probe the biophysical properties of MUT-16 and discover that Mutator foci are liquid-like condensates which assemble by phase separation. Mutator foci are adjacent to two additional phase-separated condensates: P granules, which interact with newly exported RNA, and Z granules, which are necessary for siRNA inheritance. Building on previous studies, my inaugural work proposes that RNA silencing is facilitated by multiple phase-separated condensates at the nuclear periphery of germ cells. ❧ Next, I investigate the regulation of Mutator foci and define the spatiotemporal appearance and localization of Mutator foci. I discover that Mutator foci arise in early embryos, persist in all larval stages, and are present in spermatids, indicating a role in paternal inheritance of small RNAs. I additionally demonstrate that RNA and the germline cell cycle influence the morphology of Mutator foci and suggest that these factors may influence the efficacy of RNA silencing in certain cellular environments. ❧ Lastly, I probe the interaction of Mutator foci with adjacent perinuclear compartments involved in small RNA pathways, which I refer to as C. elegans nuage. I demonstrate that nuage interaction is independent of the germline environment and able to be reestablished after disruption. I interrogate the spatial configuration of nuage compartments with superresolution microscopy techniques and discover a previously undescribed toroidal P granule morphology, in which some P granules surround Mutator foci and other nuage compartments with a consistent exterior-to-interior organization. Finally, I find that Mutator foci are the scarcest nuage compartment and that nuage compartments exist in distinct populations, suggesting a functional subdivision of different nuage assemblages. My final work aims to create a more accurate model of RNA silencing through multiple phase-separated compartments. Together, my doctoral studies begin to reveal how the complex and critical small RNA pathways are organized within C. elegans nuage.

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Asset Metadata

Creator Uebel, Celja Jae Churches (author)

Core Title The organization of small RNA pathways within C. elegans germ granules: mutator foci formation, regulation, and interaction

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Degree Conferral Date 2021-12

Publication Date 10/14/2021

Defense Date 08/05/2021

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

Tag C. elegans,germ granules,OAI-PMH Harvest,phase separation,RNA interference,small RNAs

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Phillips, Carolyn M. (committee chair), Curran, Sean P. (committee member), Pinaud, Fabien F. (committee member), Pratt, Matthew R. (committee member)

Creator Email celjajae@gmail.com,uebel@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-oUC16208290

Unique identifier UC16208290

Legacy Identifier etd-UebelCelja-10164

Document Type Dissertation

Rights Uebel, Celja Jae Churches

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

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