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SIMR-1 facilitates robust silencing of piRNA target loci in the C. elegans germline

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SIMR-1 facilitates robust silencing of piRNA target loci in the C. elegans germline

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Content SIMR-1 facilitates robust silencing of piRNA target loci in the
C. elegans germline
By
Kevin I. Manage
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
Copyright 2021 Kevin I. Manage
ii
Acknowledgments
It has been a long road but I am happy I get the opportunity to thank all those who
have been a part of my journey to this day. I want to start off by thanking Ms. Jen Bejarano
who helped ignite my passion for biology way back in the seventh grade. It was her
biology class in middle school that set me down the path that led here. To Dr. Rahul
Warrior and Dr. Steve Lipkin, it was during my time at UCI where I was first exposed to
real bench work and realized that working in the sciences could be something I saw for
myself as a career. To the friends I made at UCI, I will always remember the late-nights
at the science library fueled by junk food and energy drinks as well as all the subsequent
parties and stupid things we used to get up to. My years at UCI were some of the best of
my life and it was during that time that I did the most growing from a shy kid right out of
high school to a slightly more confident adult. Parita and James, my time working in
industry with you made what could have been a mundane 9-5 into a job that I looked
forward to day in and day out and contributed to my realization that a continuing career
in biotech would be something I would strive for post PhD. My friends in Los Angeles; I
may have met you all through Rachna but am happy to be able to call you all my friends
now as well. To BFFAEAE, you are my oldest and truly best friends. You have known me
through all my ups and down and to me, you are family. All of you have helped shape me
into who I am today and I can’t express how happy I am that we have stayed as close as
we have.
To my doctoral advisor, Dr. Carolyn Phillips, who accepted me into her lab as one
of her very first grad students, I will forever be grateful for your mentorship and guidance
through all these years and for molding me into a scientist. I honestly don’t think I could
iii
have asked for anyone better to help get through my time here. Your excitement and
enthusiasm for the work we all did was what made trying to find the answers to small RNA
silencing in these tiny little worms worth the long hours and seemingly endless number of
new questions to tackle. I can only say thank you for taking a chance on me way back
when.
To the members of my committee: Dr. Sean Curran, Dr. Bérénice Benayoun and
Dr. Matt Michael, you will never know the relief I felt after our first meeting as a group. Up
until then, I felt like nothing I had done to that point was enough but your assurances that
I was on track and nearing the end gave me the drive to push through towards the finish
line. I thank you for that, as well for all of your individual advice and insightful comments
over the years.
To the members of the Phillips lab, past and present, I thank you all. I am one of
the few who gets to say he has known every person who has come and gone over the
years and I am grateful for the comradery and friendship I developed with you all. I truly
think the quality of the lab environment and members is what can make or break an
individual’s PhD experience and I am lucky to have worked with some of the very best. I
know that our friendships will be lifelong and I am excited to see the paths you all take in
the future. As such, I promote you all from workplace-proximity associates…to friends.
Also, to my 2015 cohort, I am thankful to have come into this crazy experience with you
all. I don’t think I could have made it through those first few years without each of you and
I am proud of the scientists you have all become.
I also want to take this opportunity to thank the MCB department staff, both past
and present. They work behind the scenes to make sure that we as students are taken
iv
care of and that the building doesn’t fall apart. A special thank you to Katie Walker Boeck
for being someone who has helped me tremendously over all these years and who goes
above and beyond to make sure that we have everything that we need.
To my family, both those still here and those who left us too soon, your support
and love has meant the world to me. I have taken all the lessons learned from each of
you over the past 35 years and made them part of the foundation of who I am today. I
only hope that this accomplishment makes you proud and that you know this would not
have been possible without you all. To my parents, I love you both and even though I feel
like you still have no idea what I do or how this whole PhD thing works, I am grateful for
all the unconditional support. Also, to Dillon and Lorraine, I love you both even if I don’t
say it often enough, and thank you for being the best little brother and sister- ones that I
always had, but never wanted!
Lastly, I would like to thank my wife, Rachna. Having never lived in Los Angeles
proper and moving here only really knowing you, you made LA feel like home and helped
me develop a support system that kept me sane over the years. I was integrated into your
friend group as if I had always been a part of it and have developed many lifelong
friendships that I treasure. I know that my being in grad school has given you your own
fair share of headaches but you have always made me feel prioritized. Having you to
come home to at the end of those tough days helped keep me going throughout the last
6 years. Though I definitely don’t say it enough, you have been a source of strength and
love, and I don’t think I can ever truly express how important you are to me. Plus, now I
am officially going graduate as a USC Trojan- Zot Zot!!

v
Table of Contents
Acknowledgments .......................................................................................................... ii
Table of Contents ........................................................................................................... v
List of Tables ................................................................................................................ vii
List of Figures .............................................................................................................. viii
Abstract ........................................................................................................................... x
Chapter 1. Introduction .................................................................................................. 1
Small RNAs in Caenorhabditis elegans ................................................................ 1
The mutator complex is the site of secondary siRNA biogenesis ......................... 3
MUT-2 is critical to mutator complex function ....................................................... 4
Secondary siRNA production may involve 3’ end trimming ................................... 5
Tudor domain proteins play important roles in the piRNA silencing pathway ....... 6
Distinct perinuclear condensates control spatial organization and processing of
mRNA molecules ................................................................................................... 7
Significance ........................................................................................................... 8
References .......................................................................................................... 10
Chapter 2. A Tudor domain protein, SIMR-1, promotes siRNA production at
piRNA-targeted mRNAs in C. elegans ........................................................................ 15
Author Contributions ............................................................................................ 15
Abstract ............................................................................................................... 16
Introduction .......................................................................................................... 16
Results ................................................................................................................. 20
Discussion ........................................................................................................... 46
Materials and Methods ........................................................................................ 51
Acknowledgements ............................................................................................. 61
References .......................................................................................................... 62
Figures and Figure Legends ............................................................................... 73
Chapter 3. Proximity labeling to identify SIMR-1 protein interactors ...................... 90
Introduction .......................................................................................................... 90
Results ................................................................................................................. 93
Discussion ........................................................................................................... 97
Materials and Methods ........................................................................................ 99
References ........................................................................................................ 102
Figures and Figure Legends ............................................................................. 107
vi
Chater 4. Identifying RNA modifying proteins at multiple steps in the C. elegans
RNA silencing pathway .............................................................................................. 114
Introduction ........................................................................................................ 114
Results ............................................................................................................... 118
Discussion ......................................................................................................... 122
Materials and Methods ...................................................................................... 125
References ........................................................................................................ 137
Figures and Figure Legends ............................................................................. 141
Appendix ..................................................................................................................... 151
SIMR foci are found in the progenitor germ cells of C. elegans embryos ......... 151

vii
List of Tables
Chapter 3. Proximity Labeling to identify SIMR-1 protein interactors
Supplementary file 1. Yeast two-hybrid of SIMR-1 identified three potential
interactors. ......................................................................................................... 111
Supplementary file 2. C. elegans strain used in the study. ............................... 112
Supplementary file 3. Oligonucleotides used in this study. ............................... 113
Chapter 4. Identifying RNA modifying proteins at multiple steps in the C. elegans
RNA silencing pathway
Table 1. Exiqon miRCURY LNA detection probes ............................................ 146
Supplementary file 1. 3’-5’ exonuclease candidate proteins used in this study. 148
Supplementary file 2. C. elegans strains used in this study. ............................. 149
Supplementary file 3. Oligonucleotides sequences used in this study. ............. 150

viii
List of Figures
Chapter 2. A Tudor domain protein, SIMR-1, promotes siRNA production at
piRNA-targeted mRNAs in C. elegans
Figure 1. SIMR-1 is a perinuclear-localized Tudor domain protein. ................... 73
Figure 1–figure supplement 1. Identification and localization of MUT-16-
associated proteins. ............................................................................................ 74
Figure 2. Small RNA-related phenotypes associated with deletions in MUT-16-
associated proteins. ............................................................................................ 75
Figure 2–figure supplement 1. Mutator-class small RNAs are reduced in simr-1
but not hpo-40 mutants. ...................................................................................... 76
Figure 3. simr-1 mutants have a transgenerational fertility defect at elevated
temperature. ........................................................................................................ 77
Figure 3–figure supplement 1. hpo-40 does not contribute to the progressive
sterility of simr-1 mutants. .................................................................................... 78
Figure 4. simr-1 mutants have piRNA-related defects. ....................................... 79
Figure 4–figure supplement 1. simr-1 mutants do not display defects associated
with mutants in the mutator or ERGO-1 26G-siRNA pathways. .......................... 80
Figure 5. simr-1 mutants display reduced small RNAs mapping to mutator and
piRNA-target genes. ............................................................................................ 81
Figure 5–figure supplement 1. Small RNAs are reduced at many mutator, piRNA,
and ERGO-1 target genes in simr-1 mutants at 25°C. ........................................ 83
Figure 6. simr-1 mutants display reduced small RNAs mapping to piRNA-
dependent transposons and increased small RNAs mapping to histone genes. 84
Figure 6–figure supplement 1. Small RNAs mapping to piRNA target transposons
are reduced and small RNAs mapping to histone genes are increased in simr-1
mutants. ............................................................................................................... 86
Figure 7. SIMR-1 localizes to foci adjacent to P granules and Mutator foci. ....... 88
Figure 7–figure supplement 1. SIMR-1 and PRG-1 localize independently and to
distinct granules. ................................................................................................. 89
Chapter 3. Proximity labeling to identify SIMR-1 protein interactors
Figure 1. Proximity labeling with TurboID identifies protein-protein interactors by
biotinylation. ...................................................................................................... 107
Figure 2. The AID system allows for fast, and reversible knockdown of degron
tagged proteins. ................................................................................................. 108
Figure 3. SIMR-1 localization is unchanged following fusion to the TurboID biotin
ligase. ................................................................................................................ 109
Figure 4. SIMR-1 bound to TurboID leads to biotinylation of proteins in addition to
those endogenously biotinylated. ...................................................................... 110
ix
Chapter 4. Identifying RNA modifying proteins at multiple steps in the C. elegans
RNA silencing pathway
Figure 1. RNA silencing is strengthened at the mutator complex. .................... 141
Figure 2. 3’-5’ exonuclease knockdown showed no significant change to siRNA
length. ................................................................................................................ 142
Figure 2-figure supplement 1. miRNA and piRNA probes targeting 3’-5’
exonuclease candidate small RNAs. ................................................................. 143
Figure 3. Catalytically dead MUT-2 is RNAi defective but shows no change in
localization. ........................................................................................................ 144
Figure 4. 3’ End-Seq protocol development. ..................................................... 145
Appendix
Figure 1. SIMR foci are numerous and bright in the Z2/Z3 progenitor germ cells.
........................................................................................................................... 152

x
Abstract
The piwi-interacting RNA (piRNA) pathway is an evolutionarily conserved
mechanism that plays an important role in the silencing of transposons and other germline
genes. Transposons are selfish genetic elements that when unregulated, can damage
the integrity of an organism’s genome, impacting future generations and potentially
leading to sterility. In Caenorhabditis elegans (C. elegans), regulation of piRNA target
genes is mediated by the mutator complex, which generates high levels of secondary
siRNAs for these piRNA-target loci to achieve robust silencing. How coordination between
piRNA production in P granules and mutator complex-dependent siRNA biogenesis
occurs is not well understood.
The work presented in this dissertation explores the function of SIMR-1, a Tudor
domain protein critical to the coordination between these two important steps in the piRNA
pathway. While not necessary for piRNA production, simr-1 mutants fail to produce high
levels of secondary siRNAs for many piRNA-target loci suggesting SIMR-1 acts at a step
between these two events. It also identifies SIMR-1 localization and establishes SIMR
foci as a new germ granule among other known phase separated condensates in the
perinuclear space of the C. elegans germline. This work also develops a unique
application of proximity-dependent labeling in conjunction with the auxin-inducible
degradation system by which to identify previously undiscovered direct interactors of
SIMR-1, as well as discover additional SIMR foci components.

1
Chapter I
Introduction
Small RNAs play important roles in maintaining proper gene expression and
protecting the genome from endogenous and exogenous sources of deleterious RNA,
such as viruses and transposons. Additionally, small RNA pathways are necessary to
ensure proper development, chromosome segregation and gamete production. In
Caenorhabditis elegans (C. elegans), small RNAs can range in size from 18-30
nucleotides in length and consist of three major subclasses; microRNAs (miRNAs), piwi-
interacting RNAs (piRNAs), and small interfering RNAs (siRNAs)(Ghildiyal and Zamore,
2009).

Small RNAs in Caenorhabditis elegans
miRNAs are found in many phyla and are generally involved in downregulating
gene expression of processes such as cell fate specification, apoptosis, and metabolism
(Bartel, 2004). In C. elegans they are transcribed by RNA polymerase II as poly-
adenylated primary miRNAs that are approximately 1 kilobase in length (Vella and Slack,
2005). These molecules are then processed by the RNAse III endonuclease Drosha into
precursor miRNA that are 60-70 nucleotides and able to form hairpin loop structures. The
precursor miRNA is then processed again in the cytoplasm by an RNAse III enzyme Dicer,
that processes the precursor miRNA into mature 20-25 nucleotide miRNA (Vella and
Slack, 2005). These mature miRNAs bind to imperfectly complementary sequences in 3’
untranslated regions of target mRNAs, and begin silencing after complexing with
Argonaute effector proteins (Vella and Slack, 2005).
2
piRNAs, also known as 21U RNAs in C. elegans, are necessary to maintain fertility
and germ cell function by silencing transposons and other germline mRNAs, and are
characterized by a length of 21 nucleotides and a 5’ uridine bias (Ketting, 2011; Weick
and Miska, 2014). 21U RNAs are not transcribed by RNA dependent RNA polymerase
complexes, nor do they rely on Dicer or other factors present in the biogenesis of other
small RNAs (Batista et al., 2008; Das et al., 2008). Instead, 21U RNAs are encoded as
independent transcriptional units by Pol II and once transcribed, are bound by PRG-1, a
single functional homolog of a subgroup of Argonaute proteins called Piwi proteins
(Batista et al., 2008; Billi et al., 2013; Das et al., 2008; Gu et al., 2012; Wang and Reinke,
2008). In order to increase the effectiveness of silencing at piRNA targets, C. elegans
utilize a small RNA amplification pathway centered around the mutator complex to
generate secondary downstream siRNAs from piRNA-targeted mRNAs to trigger robust
and heritable silencing of piRNA specific targets (Ashe et al., 2012; Bagijn et al., 2012;
Das et al., 2008; Lee et al., 2012; Shirayama et al., 2012)
siRNAs can be generated from exogenous and endogenous sources of double-
stranded RNA, with primary siRNAs requiring the RNAse III-like enzyme Dicer while
secondary siRNAs utilize the mutator complex (Bernstein et al., 2001; Ketting et al., 2001;
Phillips et al., 2012). The mechanisms for identification of RNA to be silenced can vary,
but one such mechanism is the recognition of dsRNA by Dicer (Duchaine et al., 2006;
Ketting et al., 2001; Phillips et al., 2012). Upon recognition, Dicer cleaves dsRNA,
generating primary siRNAs available for binding which act as a guide for the Argonaute
protein RDE-1 to silence complementary mRNA transcripts (Carmell et al., 2002). siRNAs
are generally classified as either 26G or 22G, based on their length and 5’ nucleotide and
3
silence a variety of targets including exogenous RNA, aberrant transcripts, transposons
and pseudogenes (Gu et al., 2009; Zhang et al., 2011). 26G siRNAs are considered
primary siRNAs and are present in relatively low levels that in addition to silencing target
mRNAs, can also trigger formation of secondary 22G siRNAs at the mutator complex,
though a large number of 22G siRNAs do not require a 26G siRNA trigger (Gent et al.,
2010; Gu et al., 2009; Pak and Fire, 2007; Yigit et al., 2006; Zhang et al., 2011). These
newly amplified secondary siRNAs can be bound by worm-specific Argonaute (WAGO)
proteins downstream of the amplification step to continue silencing target mRNA. As with
the small RNA amplification that occurs in piRNAs, these secondary siRNAs are
necessary for robust, effective silencing.

The mutator complex is the site of secondary siRNA biogenesis
In organisms such as C. elegans, plants and fungi, this secondary siRNA
amplification occurs through the activity of an RNA-dependent RNA polymerase (RdRP)
as part of the mutator complex, which generates large numbers of antisense secondary
RNAs that are utilized by Argonaute effector proteins to enhance RNA silencing (Gu et
al., 2009; Phillips et al., 2012; Vasale et al., 2010). The mutator complex consists of a
collection of proteins involved in secondary siRNA biogenesis, centered around a
scaffolding protein MUT-16. MUT-16 recruits other proteins involved in amplification,
including the nucleotidyltransferase MUT-2, the 3’-5’ exonuclease MUT-7, the DEAD-box
RNA helicases MUT-14 and SMUT-1, and two uncharacterized proteins, RDE-2 and
MUT-15 (Phillips et al., 2012, 2014; Uebel et al., 2018). There are several other
components known to localize to the mutator complex but are not dependent on MUT-16
4
for their localization, including the RdRP RRF-1 (Phillips et al., 2012; Tsai et al., 2015).
The binding of these secondary siRNAs to Argonaute effector proteins allows for silencing
of complementary mRNA molecules through a variety of methods such as transcription
repression, translation inhibition and mRNA decay in an evolutionarily conserved process
called RNA interference (RNAi)(Claycomb, 2014; Hutvagner and Simard, 2008; Ketting,
2011; Ketting et al., 2001). Though much is known about the protein components of the
mutator complex, there is still a great deal to be understood about their roles and
mechanisms. One of these proteins that has been shown to be required for production of
most siRNAs, but is not completely characterized is MUT-2 (Chen et al., 2005; Gu et al.,
2009; Phillips et al., 2012; Zhang et al., 2011).

MUT-2 is critical to mutator complex function
MUT-2, based on its amino acid sequence, is predicted to be a homolog of the b-
nucleotidyltransferase family, many of which modify endo-siRNAs, miRNAs, or miRNA
precursors (Chen et al., 2005; Heo et al., 2009; Wolfswinkel et al., 2009). Proteins in this
superfamily contain a nucleotidyltransferase domain that consists of a conserved glycine-
serine motif and aspartic-acid triad. This aspartic-acid triad coordinates two divalent metal
cations involved in a two-metal ion mechanism of nucleotide addition (Martin et al., 2004).
The nucleotidyltransferase domain is commonly found in poly (A) polymerases and
terminal uridyl transferases. Mutations compromising these two features leads to an RNAi
deficient phenotype, and those same residues are conserved in the fission yeast homolog
Cid12, which associates with RdRPs and is required for S. pombe siRNA accumulation
(Chen et al., 2005). MUT-2 has recently been shown to be involved in adding stretches
5
of non-templated uridine and guanosine ribonucleotides to the 3’ ends of RNA targeted
during transposon silencing and transgenerational epigenetic inheritance (Shukla et al.,
2020). These poly(UG) or pUG tails are added to germline and soma expressed RNAs
and it has been suggested that these pUG tails help to stabilize mRNA fragments and
possibly form a structure essential for RdRP recruitment. During RNAi induced silencing,
these pUG RNAs persist for several generations, but are not permanent. Over repeated
cycles of siRNA synthesis, pUG RNAs shorten during each round, possibly as a result of
RdRP-dependent secondary siRNA synthesis. However, the natural targets of pUG tailing
such as transposons, are silenced indefinitely, suggesting that endogenous targets of
pUG tailing are reinforced at each generation (Shukla et al., 2020).

Secondary siRNA production may involve 3’ end trimming
The mechanism of biogenesis of secondary siRNAs and what factors determine
their exact length of 22 nucleotides is not fully understood. It is possible that the
processivity of the RdRP RRF-1 is sufficient to generate siRNAs of precisely 22
nucleotides, but it is more likely that a 3’-5’ exonuclease contributes to the generation of
correctly-sized siRNAs. This idea is supported by evidence of 3’ end trimming in a variety
of small RNA pathways, in numerous organisms. Members of the PARN 3’-5’
exonuclease family, PARN-1 and PNLDC1 in C. elegans and B. mori respectively, have
been found to trim the 3’ ends of piRNAs while Nibbler, a 3’-5’ exoribonuclease in D.
melanogaster, is involved in miRNA 3’ end trimming (Han et al., 2011; Izumi et al., 2016;
Tang et al., 2016). These findings suggest that despite differences between small RNA
types and between species, the mechanisms for small RNA trimming may be conserved.
6
Without proper processing, amplification of the silencing signal will be lost as WAGO
proteins have specific binding requirements with regard to transcript length.

Tudor domain proteins play important roles in the piRNA silencing pathway
The piRNA pathway is critical to normal germline development, specifically in the
regulation of transposons and other germline genes, and the components of this pathway
are primarily expressed in the C. elegans germline (Das et al., 2008). In addition to 21U
RNAs and the Argonaute protein PRG-1, another major component of the piRNA pathway
is the Tudor domain protein family. Tudor domain proteins play important roles in piRNA
accumulation and mRNA target regulation through their PIWI protein interactions in
animals such as mice and Drosophila (Chen et al., 2011; Nishida et al., 2009; Reuter et
al., 2009). Tudor domains play a role in facilitating protein-protein interactions in a
methylation specific manner by recognizing methylated arginines or lysines (Chen et al.,
2011; Friesen et al., 2001). However, Tudor domain proteins that are affiliated with the
piRNA pathway often interact with an additional conserved element flanking the Tudor
domain core referred to as the extended Tudor domain. The extended Tudor domain is
necessary for the Tudor domain protein’s ability to recognize peptides containing a
methylated arginine (Chen et al., 2011; Liu et al., 2010a, 2010b). The extended Tudor
domain preferentially recognizes symmetrically dimethylated arginine (sDMA)
modifications over monomethylated arginines (MMA), asymmetrically dimethylated
arginines (aDMA) or non-methylated peptides, though some extended Tudor domain
proteins have lost the ability to bind methylated arginine and can only recognize
unmodified peptides (Liu et al., 2010b; Zhang et al., 2017).
7
Distinct perinuclear condensates control spatial organization and processing of
mRNA molecules
Many components of the various small RNA silencing pathways, including those
components in the piRNA pathway, are localized to perinuclear, membrane-less
compartments surrounding germline nuclei. It has been shown that these, as well other
P granule components assemble by intracellular phase separation (Brangwynne et al.,
2009). Mutator foci, sites of secondary siRNA biogenesis by the mutator complex, and Z
granules, required for RNAi inheritance, have also shown to be phase-separated
biomolecular condensates that lie next to each other, and P granules at the nuclear
periphery (Uebel et al., 2018; Wan et al., 2018).
In Drosophila, components of the piRNA pathway including PIWI and Tudor
domain proteins localize to a compartment known as nuage, while in C. elegans the
proteins localize to an analogous structure referred to as the P granule. P granules are
known to have over 50 protein components and are closely associated with nuclear pores
in the C. elegans germline, suggesting that mRNAs leaving the nucleus pass through
them (Sundby et al., 2021). P granules have many roles including germ cell fate
determination, transmitting epigenetic information, and post-transcriptional gene
regulation (Ashe et al., 2012; Batista et al., 2008; Shirayama et al., 2012).
Z granules are perinuclear foci with only three currently known components, ZNFX-
1, WAGO-4 and PID-2. These foci are thought to form in the region between P granules
and Mutator foci in the perinuclear space (Wan et al., 2018). The main role Z granules
appear to play is in marking mRNA for heritable silencing and triggering transgenerational
epigenetic inheritance (Ishidate et al., 2018; Lev et al., 2019; Wan et al., 2018). This
8
grouping of condensates has come to be referred to as PZM granules or nuage. The
discovery of this collection of membrane-less condensates has led to a model where the
small RNA pathway is temporally and spatially organized, with each step of the silencing
pathway occurring in distinct condensates while still allowing for RNAs and perhaps some
proteins to be trafficked between them.

Significance
Though there has been a great deal of research conducted in C. elegans over the
years regarding the various, complicated pathways that fall under the umbrella of small
RNA silencing, there is much that is still not well understood. When I began my thesis
project I sought to identify and characterize novel components of small RNA silencing
pathways.
In my second chapter, I have characterized a previously unknown protein, SIMR-
1 (siRNA-defective and mortal germline). SIMR-1 is a Tudor domain protein that
associates with the core component of the mutator complex, MUT-16, but has a
phenotype more similar to that of piRNA pathway mutants. SIMR-1 forms perinuclear foci
adjacent to, but distinct from previously characterized granules and first appear during
the P4 blastomere, becoming numerous and bright in the Z2 and Z3 progenitor germ cells
(see Appendix). Through sequencing analysis, I was able to demonstrate that SIMR-1
acts in the piRNA pathway, at a step downstream of PRG-1 targeting and plays an
important role in promoting siRNA biogenesis at piRNA targets by the mutator complex.
In my third chapter, I sought to develop a proximity-dependent labeling technique
for use in the C. elegans germline, specifically in the identification of SIMR-1 protein
9
interactors. Through this work, I have generated SIMR-1 fused to a promiscuous biotin
ligase that can biotinylate neighboring proteins for mass spectrometry-based protein
identification. To combat the issue of high levels of endogenous biotinylated proteins, I
tagged, BPL-1, a C. elegans biotinylase involved in fatty acid biosynthesis, with an auxin-
inducible degradation (AID) tag for tissue-specific and temporal knockdown (Watts et al.,
2018; Zhang et al., 2015). Use of these two systems together will allow for depletion of
endogenous biotinylated proteins, while simultaneously biotinylating SIMR-1 interacting
and neighboring proteins, with the ultimate goal of purification of these biotinylated
proteins by streptavidin pulldown and identification by mass spectrometry.
In my fourth chapter, I focus on the role of exonucleases and nucleotidyl
transferases in RNA silencing. First, I attempted to identify the 3’-5’ exonuclease that trims
newly transcribed secondary siRNAs to their appropriate 22-nucleotide length. I then
examined the role of MUT-2 in the modification of mRNA templates to be used for
secondary siRNA biogenesis. The completion of this work will elucidate how the proteins
involved in making modifications to both small RNAs and mRNAs in C. elegans are
necessary to ensuring robust RNA silencing.
Together, my doctoral work has identified novel components of the RNA silencing
pathway and discovered a new type of germ granule. Through this research, I have
contributed a small piece towards the further understanding of the large and complicated
field of C. elegans small RNA silencing, which can be built upon for future discoveries.

10
References
Ashe, A., Sapetschnig, A., Weick, E.-M., Mitchell, J., Bagijn, M.P., Cording, A.C.,
Doebley, A.-L., Goldstein, L.D., Lehrbach, N.J., Pen, J.L., et al. (2012). piRNAs Can
Trigger a Multigenerational Epigenetic Memory in the Germline of C. elegans. Cell
150, 88–99.
Bagijn, M.P., Goldstein, L.D., Sapetschnig, A., Weick, E.-M., Bouasker, S., Lehrbach,
N.J., Simard, M.J., and Miska, E.A. (2012). Function, targets, and evolution of
Caenorhabditis elegans piRNAs. Science 337, 574–578.
Bartel, D.P. (2004). MicroRNAs Genomics, Biogenesis, Mechanism, and Function. Cell
116, 281–297.
Batista, P.J., Ruby, J.G., Claycomb, J.M., Chiang, R., Fahlgren, N., Kasschau, K.D.,
Chaves, D.A., Gu, W., Vasale, J.J., Duan, S., et al. (2008). PRG-1 and 21U-RNAs Interact
to Form the piRNA Complex Required for Fertility in C. elegans. Mol Cell 31, 67–78.
Bernstein, E., Caudy, A.A., Hammond, S.M., and Hannon, G.J. (2001). Role for a
bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363–366.
Billi, A.C., Freeberg, M.A., Day, A.M., Chun, S.Y., Khivansara, V., and Kim, J.K. (2013).
A Conserved Upstream Motif Orchestrates Autonomous, Germline-Enriched Expression
of Caenorhabditis elegans piRNAs. Plos Genet 9, e1003392.
Brangwynne, C.P., Eckmann, C.R., Courson, D.S., Rybarska, A., Hoege, C., Gharakhani,
J., Jülicher, F., and Hyman, A.A. (2009). Germline P Granules Are Liquid Droplets That
Localize by Controlled Dissolution/Condensation. Science 324, 1729–1732.
Carmell, M.A., Xuan, Z., Zhang, M.Q., and Hannon, G.J. (2002). The Argonaute family:
tentacles that reach into RNAi, developmental control, stem cell maintenance, and
tumorigenesis. Genes & Development 16, 2733–2742.
Chen, C., Nott, T.J., Jin, J., and Pawson, T. (2011). Deciphering arginine methylation:
Tudor tells the tale. Nat Rev Mol Cell Bio 12, 629–642.
Chen, C.-C.G., Simard, M.J., Tabara, H., Brownell, D.R., McCollough, J.A., and Mello,
C.C. (2005). A Member of the Polymerase β Nucleotidyltransferase Superfamily Is
Required for RNA Interference in C. elegans. Current Biology 15, 378–383.
Claycomb, J.M. (2014). Ancient Endo-siRNA Pathways Reveal New Tricks. 24, R703–
R715.

11
Das, P.P., Bagijn, M.P., Goldstein, L.D., Woolford, J.R., Lehrbach, N.J., Sapetschnig, A.,
Buhecha, H.R., Gilchrist, M.J., Howe, K.L., Stark, R., et al. (2008). Piwi and piRNAs Act
Upstream of an Endogenous siRNA Pathway to Suppress Tc3 Transposon Mobility in the
Caenorhabditis elegans Germline. Mol Cell 31, 79–90.
Duchaine, T.F., Wohlschlegel, J.A., Kennedy, S., Bei, Y., Jr, D.C., Pang, K., Brownell,
D.R., Harding, S., Mitani, S., Ruvkun, G., et al. (2006). Functional Proteomics Reveals
the Biochemical Niche of C. elegans DCR-1 in Multiple Small-RNA-Mediated Pathways.
Cell 124, 343–354.
Friesen, W.J., Massenet, S., Paushkin, S., Wyce, A., and Dreyfuss, G. (2001). SMN, the
Product of the Spinal Muscular Atrophy Gene, Binds Preferentially to Dimethylarginine-
Containing Protein Targets. Mol Cell 7, 1111–1117.
Gent, J.I., Lamm, A.T., Pavelec, D.M., Maniar, J.M., Parameswaran, P., Tao, L.,
Kennedy, S., and Fire, A.Z. (2010). Distinct Phases of siRNA Synthesis in an Endogenous
RNAi Pathway in C. elegans Soma. Mol Cell 37, 679–689.
Ghildiyal, M., and Zamore, P.D. (2009). Small silencing RNAs: an expanding universe.
Nature Reviews. Genetics 10, 94–108.
Gu, W., Shirayama, M., Conte, D., Vasale, J., Batista, P.J., Claycomb, J.M., Moresco,
J.J., Youngman, E.M., Keys, J., Stoltz, M.J., et al. (2009). Distinct argonaute-mediated
22G-RNA pathways direct genome surveillance in the C. elegans germline. Molecular
Cell 36, 231–244.
Gu, W., Lee, H.-C., Chaves, D., Youngman, E.M., Pazour, G.J., Jr, D.C., and Mello, C.C.
(2012). CapSeq and CIP-TAP Identify Pol II Start Sites and Reveal Capped Small RNAs
as C. elegans piRNA Precursors. 151, 1488–1500.
Han, B.W., Hung, J.-H., Weng, Z., Zamore, P.D., and Ameres, S.L. (2011). The 3’-to-5’
exoribonuclease Nibbler shapes the 3’ ends of microRNAs bound to Drosophila
Argonaute1. Current Biology : CB 21, 1878–1887.
Heo, I., Joo, C., Kim, Y.-K., Ha, M., Yoon, M.-J., Cho, J., Yeom, K.-H., Han, J., and Kim,
V.N. (2009). TUT4 in Concert with Lin28 Suppresses MicroRNA Biogenesis through Pre-
MicroRNA Uridylation. Cell 138, 696–708.
Hutvagner, G., and Simard, M.J. (2008). Argonaute proteins: key players in RNA
silencing. Nature Reviews Molecular Cell Biology 9, 22–32.
Ishidate, T., Ozturk, A.R., Durning, D.J., Sharma, R., Shen, E.-Z., Chen, H., Seth, M.,
Shirayama, M., and Mello, C.C. (2018). ZNFX-1 Functions within Perinuclear Nuage to
Balance Epigenetic Signals. 70, 639-649.e6.
12
Izumi, N., Shoji, K., Sakaguchi, Y., Honda, S., Kirino, Y., Suzuki, T., Katsuma, S., and
Tomari, Y. (2016). Identification and Functional Analysis of the Pre-piRNA 3′ Trimmer in
Silkworms. Cell 164, 962–973.
Ketting, R.F. (2011). The Many Faces of RNAi. Developmental Cell 20, 148–161.
Ketting, R.F., Fischer, S.E., Bernstein, E., Sijen, T., Hannon, G.J., and Plasterk, R.H.
(2001). Dicer functions in RNA interference and in synthesis of small RNA involved in
developmental timing in C. elegans. Genes & Development 15, 2654–2659.
Lee, H.-C., Gu, W., Shirayama, M., Youngman, E., Conte, D., and Mello, C.C. (2012).
C. elegans piRNAs Mediate the Genome-wide Surveillance of Germline Transcripts. Cell
150, 78–87.
Lev, I., Toker, I.A., Mor, Y., Nitzan, A., Weintraub, G., Antonova, O., Bhonkar, O.,
Shushan, I.B., Seroussi, U., Claycomb, J.M., et al. (2019). Germ Granules Govern Small
RNA Inheritance. Current Biology 1–28.
Liu, H., Wang, J.-Y.S., Huang, Y., Li, Z., Gong, W., Lehmann, R., and Xu, R.-M. (2010a).
Structural basis for methylarginine-dependent recognition of Aubergine by Tudor. Gene
Dev 24, 1876–1881.
Liu, K., Chen, C., Guo, Y., Lam, R., Bian, C., Xu, C., Zhao, D.Y., Jin, J., MacKenzie, F.,
Pawson, T., et al. (2010b). Structural basis for recognition of arginine methylated Piwi
proteins by the extended Tudor domain. Proc National Acad Sci 107, 18398–18403.
Martin, G., Möglich, A., Keller, W., and Doublié, S. (2004). Biochemical and Structural
Insights into Substrate Binding and Catalytic Mechanism of Mammalian Poly(A)
Polymerase. Journal of Molecular Biology 341, 911–925.
Nishida, K.M., Okada, T.N., Kawamura, T., Mituyama, T., Kawamura, Y., Inagaki, S.,
Huang, H., Chen, D., Kodama, T., Siomi, H., et al. (2009). Functional involvement of
Tudor and dPRMT5 in the piRNA processing pathway in Drosophila germlines. Embo J
28, 3820–3831.
Pak, J., and Fire, A. (2007). Distinct populations of primary and secondary effectors
during RNAi in C. elegans. Science 315, 241–244.
Phillips, C.M., Montgomery, T.A., Breen, P.C., and Ruvkun, G. (2012). MUT-16 promotes
formation of perinuclear mutator foci required for RNA silencing in the C. elegans
germline. Genes & Development 26, 1433–1444.
Phillips, C.M., Montgomery, B.E., Breen, P.C., Roovers, E.F., Rim, Y.-S., Ohsumi, T.K.,
Newman, M.A., Wolfswinkel, J.C. van, Ketting, R.F., Ruvkun, G., et al. (2014). MUT-14
and SMUT-1 DEAD Box RNA Helicases Have Overlapping Roles in Germline RNAi and
Endogenous siRNA Formation. Current Biology 24, 839–844.
13
Reuter, M., Chuma, S., Tanaka, T., Franz, T., Stark, A., and Pillai, R.S. (2009). Loss of
the Mili-interacting Tudor domain–containing protein-1 activates transposons and alters
the Mili-associated small RNA profile. Nat Struct Mol Biol 16, 639–646.
Shirayama, M., Seth, M., Lee, H.-C., Gu, W., Ishidate, T., Jr, D.C., and Mello, C.C. (2012).
piRNAs Initiate an Epigenetic Memory of Nonself RNA in the C. elegans Germline.
Cell 150, 65–77.
Shukla, A., Yan, J., Pagano, D.J., Dodson, A.E., Fei, Y., Gorham, J., Seidman, J.G.,
Wickens, M., and Kennedy, S. (2020). poly(UG)-tailed RNAs in genome protection and
epigenetic inheritance. Nature 1–24.
Sundby, A.E., Molnar, R.I., and Claycomb, J.M. (2021). Connecting the Dots: Linking
Caenorhabditis elegans Small RNA Pathways and Germ Granules. Trends Cell Biol 31,
387–401.
Tang, W., Tu, S., Lee, H.-C., Weng, Z., and Mello, C.C. (2016). The RNase PARN-1 Trims
piRNA 3’ Ends to Promote Transcriptome Surveillance in C. elegans. Cell 164, 974–984.
Tsai, H.-Y., Chen, C.-C.G., Jr, D.C., Moresco, J.J., Chaves, D.A., Mitani, S., III, J.R.Y.,
Tsai, M.-D., and Mello, C.C. (2015). A Ribonuclease Coordinates siRNA Amplification
and mRNA Cleavage during RNAi. Cell 160, 407–419.
Uebel, C.J., Anderson, D.C., Mandarino, L.M., Manage, K.I., Aynaszyan, S., and Phillips,
C.M. (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 Genetics 14, e1007542-22.
Vasale, J.J., Gu, W., Thivierge, C., Batista, P.J., Claycomb, J.M., Youngman, E.M.,
Duchaine, T.F., Mello, C.C., and Conte, D. (2010). Sequential rounds of RNA-dependent
RNA transcription drive endogenous small-RNA biogenesis in the ERGO-1/Argonaute
pathway. Proceedings of the National Academy of Sciences of the United States of
America 107, 3582–3587.
Vella, M.C., and Slack, F.J. (2005). C. elegans microRNAs. WormBook : The Online
Review of C. Elegans Biology 1–9.
Wan, G., Fields, B.D., Spracklin, G., Shukla, A., Phillips, C.M., and Kennedy, S. (2018).
Spatiotemporal regulation of liquid-like condensates in epigenetic inheritance. Nature 1–
23.
Wang, G., and Reinke, V. (2008). A C. elegans Piwi, PRG-1, Regulates 21U-RNAs during
Spermatogenesis. Curr Biol 18, 861–867.
14
Watts, J.S., Morton, D.G., Kemphues, K.J., and Watts, J.L. (2018). The biotin-ligating
protein BPL-1 is critical for lipid biosynthesis and polarization of the Caenorhabditis
elegansembryo. The Journal of Biological Chemistry 293, 610–622.
Weick, E.-M., and Miska, E.A. (2014). piRNAs: from biogenesis to function. Development
(Cambridge, England) 141, 3458–3471.
Wolfswinkel, J.C. van, Claycomb, J.M., Batista, P.J., Mello, C.C., Berezikov, E., and
Ketting, R.F. (2009). CDE-1 Affects Chromosome Segregation through Uridylation of
CSR-1-Bound siRNAs. Cell 139, 135–148.
Yigit, E., Batista, P.J., Bei, Y., Pang, K.M., Chen, C.-C.G., Tolia, N.H., Joshua-Tor, L.,
Mitani, S., Simard, M.J., and Mello, C.C. (2006). Analysis of the C. elegans Argonaute
Family Reveals that Distinct Argonautes Act Sequentially during RNAi. Cell 127, 747–
757.
Zhang, C., Montgomery, T.A., Gabel, H.W., Fischer, S.E.J., Phillips, C.M., Fahlgren, N.,
Sullivan, C.M., Carrington, J.C., and Ruvkun, G. (2011). mut-16 and other mutator class
genes modulate 22G and 26G siRNA pathways in Caenorhabditis elegans. Proceedings
of the National Academy of Sciences of the United States of America 108, 1201–1208.
Zhang, H., Liu, K., Izumi, N., Huang, H., Ding, D., Ni, Z., Sidhu, S.S., Chen, C., Tomari,
Y., and Min, J. (2017). Structural basis for arginine methylation-independent recognition
of PIWIL1 by TDRD2. Proc National Acad Sci 114, 12483–12488.
Zhang, L., Ward, J.D., Cheng, Z., and Development, A.D. (2015). The auxin-inducible
degradation (AID) system enables versatile conditional protein depletion in C. elegans.
Dev.Biologists.Org.

15
Chapter II
A Tudor domain protein, SIMR-1, promotes siRNA production at
piRNA-targeted mRNAs in C. elegans

This manuscript was first published as:
Manage, K.I., Rogers, A.K., Wallis, D.C., Uebel, C.J., Anderson, D.C., Nguyen, D.A.H.,
Arca, K., Brown, K.C., Rodrigues, R.J.C., Albuquerque, B.F.M. de, et al. (2020). A Tudor
domain protein, SIMR-1, promotes siRNA production at piRNA-targeted mRNAs in C.
elegans. ELife 9, 1–33.

Author Contributions
A majority of the research I conducted during my time in the Phillips lab was
focused around a protein called C06A5.6, which I later renamed SIMR-1 (siRNA-
defective and mortal germline). SIMR-1 was first identified , along with several other
uncharacterized proteins, through a co-immunoprecipitation of MUT-16 by Dr. Carolyn
Phillips during her post-doctoral work. With the start of the Phillips lab at USC, Dr. Dorian
Anderson generated strains containing fluorescently tagged or mutant versions of these
proteins. It was at this time that I joined the lab and developed a research plan to begin
characterization of each protein, and in particular, SIMR-1 and its role in the piRNA
pathway.
Over the course of my investigation into the specific role of SIMR-1 in RNA
silencing, I received assistance from members of the Montgomery lab as well as Dr.
Carolyn Phillips and Dr. Alicia Rogers with analysis of sequencing data I generated.
During the final phase of completing SIMR-1 characterization, the Ketting lab contributed
reagents necessary to visualize the effects of SIMR-1 loss in the piRNA pathway. During
16
the entirety of my work on SIMR-1, I was involved in all investigation as well as
conceptualization, formal analysis and the writing of this manuscript.

Abstract
piRNAs play a critical role in the regulation of transposons and other germline
genes. In Caenorhabditis elegans, regulation of piRNA target genes is mediated by the
mutator complex, which synthesizes high levels of siRNAs through the activity of an RNA-
dependent RNA polymerase. However, the steps between mRNA recognition by the
piRNA pathway and siRNA amplification by the mutator complex are unknown. Here, we
identify the Tudor domain protein, SIMR-1, as acting downstream of piRNA production
and upstream of mutator complex-dependent siRNA biogenesis. Interestingly, SIMR-1
also localizes to distinct subcellular foci adjacent to P granules and Mutator foci, two
phase-separated condensates that are the sites of piRNA-dependent mRNA recognition
and mutator complex-dependent siRNA amplification, respectively. Thus, our data
suggests a role for multiple perinuclear condensates in organizing the piRNA pathway
and promoting mRNA regulation by the mutator complex.

Introduction
In many eukaryotes, small RNAs, ranging from ~18-30 nucleotides in length, regulate
cellular mRNAs through sequence complementarity. Argonaute proteins are key
mediators of RNA silencing; by binding to small RNAs, which interact with fully or partially
complementary mRNAs, the Argonaute proteins can promote transcription repression,
translation inhibition, and RNA decay of targeted mRNAs (Hutvagner and Simard 2008;
17
Claycomb 2014). Through this regulation of both endogenous and foreign RNAs, small
RNAs play key roles in maintaining proper gene expression and silencing deleterious
RNAs (Claycomb 2014; Ketting 2011).
A subclass of small RNAs, known as piRNAs, is critical for germ cell function, including
silencing of transposons and other germline mRNAs (Ketting 2011; Weick and Miska
2014). piRNAs are bound by a subgroup of Argonaute proteins called Piwi proteins, of
which C. elegans has a single functional homolog, PRG-1 (Batista et al. 2008; Das et al.
2008; Wang and Reinke 2008). In many organisms, including mammals, flies, and
zebrafish, piRNAs are amplified through the ping-pong mechanism (Brennecke et al.
2007; Aravin et al. 2007; Gunawardane et al. 2007; Houwing et al. 2007). This
mechanism, however, is not found in nematodes. Rather, C. elegans employs a different
mechanism to reinforce silencing at piRNA target loci. In C. elegans, a small RNA
amplification pathway dependent on the mutator complex, which includes an RNA-
dependent RNA polymerase, synthesizes secondary downstream siRNAs from piRNA-
targeted mRNAs to trigger robust and heritable silencing (Das et al. 2008; Lee et al. 2012;
Bagijn et al. 2012; Shirayama et al. 2012; Ashe et al. 2012). These siRNAs are
approximately 22-nt long, often start with a 5’G, and are bound by the WAGO clade of
Argonaute proteins, including WAGO-1, therefore, they are often referred to as WAGO-
class 22G-siRNAs (Pak and Fire 2007; Sijen et al. 2007; Yigit et al. 2006; Gu et al. 2009).
In addition to the Piwi proteins, a major player in the piRNA pathway is the Tudor
domain protein family. Tudor domain proteins in many organisms, including both mouse
and Drosophila, play critical roles in piRNA accumulation and mRNA target regulation
through their interaction with PIWI proteins (Reuter et al. 2009; Chen et al. 2011; Nishida
18
et al. 2009). The Tudor domain is a conserved structural motif originally identified in the
Drosophila protein Tudor (Boswell and Mahowald 1985; Ponting 1997; Callebaut and
Mornon 1997). Tudor domains, which function as protein-protein interaction modules,
recognize methylated arginines or lysines and thus can mediate protein interactions in a
methylation-specific manner (Friesen et al. 2001; Chen et al. 2011). Most often,
methylarginine-binding Tudor domain proteins are associated with RNA metabolism,
while methyllysine-binding Tudor domain proteins are involved in chromatin biology
(Chen et al. 2011). Interestingly, Tudor domain proteins affiliated with the piRNA pathway
often interact with an additional conserved element flanking the Tudor domain core
referred to as the extended Tudor domain, which is required for their ability to recognize
peptides containing a methylated arginine modification (Chen et al. 2011; Liu et al. 2010a;
2010b). The extended Tudor domain preferentially recognizes symmetrically
dimethylated arginine (sDMA) modifications over monomethylated arginines (MMA),
asymmetrically dimethylated arginines (aDMA), or unmodified peptides; however, some
extended Tudor domain proteins have lost the ability to bind the methylated arginine mark
and recognize only unmodified peptides (Liu et al. 2010b; Zhang et al. 2017). These
arginine methylation modifications are often found within the context of arginine-glycine
(RG) and arginine-alanine (RA) repeats and are catalyzed by the activity of Protein
Arginine Methyl Transferases (PRMTs) (Kirino et al. 2009; Vagin et al. 2009; Reuter et al.
2009; Webster et al. 2015; Liu et al. 2010a; Nishida et al. 2009).
Many components of the piRNA pathway, including some Piwi and Tudor domain
proteins, are localized to membrane-less, cytoplasmic compartments at the periphery of
germline nuclei. In Drosophila, the piRNA pathway components localize to a compartment
19
referred to as nuage, and in C. elegans, these components localize to the P granule.
Seminal work in C. elegans has shown that P granules assemble by intracellular phase
separation (Brangwynne et al. 2009). More recently, both Mutator foci, the sites of
secondary siRNA biogenesis by the mutator complex, and Z granules, which are required
for RNAi inheritance, have been shown to be phase-separated biomolecular condensates
which lie adjacent to one another and the P granule at the nuclear periphery (Uebel et al.
2018; Wan et al. 2018). This assembly of condensates can be referred to as PZM
granules or as nuage. These discoveries have led to an intriguing model where the small
RNA pathway is temporally and spatially organized into membrane-less organelles, with
distinct steps of the silencing pathway occurring in neighboring condensates, while still
allowing for trafficking of RNAs and perhaps some proteins between condensates.
Here we identify a protein required to coordinate RNA silencing between the piRNA
pathway in P granules and siRNA amplification in Mutator foci. Specifically, through
proteomic analysis of MUT-16, we identified an uncharacterized Tudor domain protein,
SIMR-1 (siRNA-defective and mortal germline). Unlike mut-16 mutants, simr-1 mutants
are not defective in exogenous RNAi, but do have a transgenerational sterility phenotype
at elevated temperature. Interestingly, while SIMR-1 is not required for production of
piRNAs or the expression of PRG-1, simr-1 mutants fail to produce high levels of siRNAs
from many piRNA-target loci. These data suggest that SIMR-1 may act at a step in
between PRG-1 targeting and siRNA biogenesis by the mutator complex. Finally, we
demonstrate that SIMR-1 localizes to perinuclear foci, adjacent to, but distinct from
Mutator foci, P granules and Z granules, which we name SIMR foci. Therefore, this work
identifies SIMR-1 as a factor that acts downstream of PRG-1 to mediate the production
20
of secondary siRNAs by the mutator complex, and suggests a role for multiple perinuclear
condensates to promote mRNA regulation by the piRNA pathway and mutator complex.

Results
Identification of MUT-16-associated proteins by functional proteomics
Many components of the mutator complex have been identified through forward and
reverse genetic screens (Supplementary File 1, Manage et al 2020) (Ketting and Plasterk
2000; Ketting et al. 1999; Tabara et al. 1999; Vastenhouw et al. 2003). More recently,
three Zc3h12a ribonuclease-like proteins that interact with the mutator complex were
identified through co-immunoprecipitation followed by mass spectrometry (IP-mass spec)
(Tsai et al. 2015). We sought to take a similar approach and extend the list of mutator
complex proteins and proteins that interact with the mutator complex. Because MUT-16
is a scaffolding protein required for assembly of the mutator complex (Phillips et al. 2012),
we chose to use an endogenously tagged MUT-16::GFP::3xFLAG for
immunoprecipitation. Following separate immunoprecipitations with GFP and FLAG
antibodies and mass spectrometry analyses, we limited our candidate list to proteins that
were present in both MUT-16-GFP and MUT-16-FLAG immunoprecipitations and absent
in both wild-type immunoprecipitations. In total, we identified 17 candidate MUT-16
interactors, twelve of which comprise all known members of the mutator complex (Phillips
et al. 2012; Uebel et al. 2018; Tsai et al. 2015) and five previously uncharacterized
proteins (Figure 1A and Supplementary File 2, Manage et al 2020). We additionally chose
to further examine three proteins (RSD-2, WAGO-1, and MATH-33) that were present in
the MUT-16-GFP immunoprecipitation, absent in the control GFP immunoprecipitation,
21
and enriched at least four-fold in the MUT-16-FLAG immunoprecipitations relative to the
control FLAG immunoprecipitation (Figure 1-figure supplement 1A and Supplementary
File 2, Manage et al 2020). RSD-2 is a small RNA factor required for exogenous RNAi
introduced at low doses and not previously known to interact with the mutator complex
(Sakaguchi et al. 2014; Han et al. 2008; Tijsterman et al. 2004; Zhang et al. 2012);
WAGO-1 is an Argonaute protein that localizes to P granules but was found to interact
with MUT-16 in a yeast two-hybrid screen (Supplementary File 1, Manage et al 2020) (Gu
et al. 2009; Phillips et al. 2014); and MATH-33 is a ubiquitin C-terminal hydrolase that
was previously identified in a proteomics screen of RDE-10-interacting proteins and RNAi
screen for genes involved in co-suppression, a phenomenon where repetitive transgenes
silence homologous endogenous genes (Zhang et al. 2012; Robert et al. 2005).
Therefore, in total, our mass spectrometry screen identified eight proteins not previously
known to be members of the mutator complex, five of which have no known link to any
small RNA pathway.

Localization of MUT-16-associated proteins
To determine whether any of the candidate MUT-16-associated proteins have
localization patterns similar to MUT-16, we tagged each protein at its endogenous locus
with a C-terminal mCherry and 2xHA tag using CRISPR. Two of the uncharacterized
proteins, MATH-33 and Y57G11C.3 localize to the nucleus of germ cells and three more,
F37C4.5, HGO-1, and C33G3.6, showed no obvious fluorescence in the cytoplasm or
nucleus of germ cells above background levels, (Figure 1-figure supplement 1B). In
contrast, C06A5.6, which we subsequently named SIMR-1, formed distinct perinuclear
22
foci in germ cells, either adjacent to or colocalizing with Mutator foci (Figure 1B). Similarly,
RSD-2 also localized to similar perinuclear foci, in contrast to previous reports that it
localizes to germ cell nuclei or the nucleolus (Figure 1-figure supplement 1C) (Sakaguchi
et al. 2014; Han et al. 2008).
Because we could not initially identify any conserved domains in SIMR-1 that would
help to predict its function, we first investigated whether there are similar proteins in C.
elegans or other related nematode species. Using BLAST, we identified a single paralog
in C. elegans, HPO-40, and a single ortholog of both SIMR-1 and HPO-40 in C. brenneri,
C. remanei, and C. japonica. SIMR-1 and HPO-40 are more closely related to one another
than to C. brenneri, C. remanei, or C. japonica paralogs, suggesting that they may be a
recent duplication (Figure 1C). We proceeded to tag HPO-40 with a C-terminal mCherry
and 2xHA tag using CRISPR, and like SIMR-1, HPO-40 formed perinuclear foci in germ
cells, either adjacent to or colocalizing with Mutator foci (Figure 1-figure supplement 1D).
MUT-16 is required for the localization of all known mutator complex proteins to
Mutator foci (Phillips et al. 2012; Uebel et al. 2018). To determine if MUT-16 is required
for SIMR-1 localization, we crossed a mut-16 null allele into the SIMR-1::mCherry strain.
Interestingly, SIMR-1 foci were still present in the mut-16 mutant (Figure 1D). To address
the reciprocal question, whether SIMR-1 or it’s paralog HPO-40 is required for MUT-16
localization, we generated deletion alleles of both simr-1 and hpo-40 using CRISPR.
MUT-16 foci were unperturbed in the simr-1; hpo-40 double mutant (Figure 1D). These
data indicate that while SIMR-1 forms germline foci near Mutator foci, it neither requires
Mutator foci for its localization, nor is the localization of Mutator foci dependent on SIMR-
1 or HPO-40, suggesting it may form separate and distinct germline foci.
23
SIMR-1 contains an extended Tudor domain
Interestingly, while a search of the Conserved Domain Database for either SIMR-1 or
HPO-40 does not identify any conserved domains, a similar search with C. remanei
CRE08315 weakly identifies a Tudor domain near the N-terminus (E-value 1.58e-03)
(Marchler-Bauer et al. 2011). We next searched SIMR-1 and related protein sequences
using the HHpred server, which is more sensitive than BLAST in finding remote homologs
(Söding et al. 2005). HHpred identified homology to multiple Tudor domain-containing
proteins, specifically those containing extended Tudor domains, including D.
melanogaster Tudor, Papi and Tudor-SN, M. musculus TDRD1, H. sapiens TDRD1,
TDRKH, and TDRD11, and B. mori Papi (Figure 1E). Many of these hits are Tudor domain
proteins with known roles in the piRNA pathway, (Liu et al. 2010a; Mathioudakis et al.
2012; Friberg et al. 2009; Ren et al. 2014; Zhang et al. 2017; 2018b). Like canonical
Tudor domains, the extended Tudor domain has four conserved aromatic residues that
form an “aromatic cage” which mediates interaction with the methylated arginine (Liu et
al. 2010a; 2010b). SIMR-1 is missing two of these four aromatic residues, making it
unclear whether it is functional to recognize a methylated substrate (Figure 1E). It does,
however, contain the absolutely conserved arginine and aspartic acid residues, which
play a structural role in the extended Tudor domain (Liu et al. 2010a). Thus, SIMR-1 is an
extended Tudor domain protein with homology to several Piwi-binding proteins. However,
further experiments will be needed to determine whether it is functionally able to
recognize methylated substrates.

24
RNA-silencing phenotypes of MUT-16-associated proteins
If any of the previously uncharacterized proteins identified in the MUT-16 IP-mass
spectrometry experiment play a role in RNA silencing, we would expect them to have
phenotypes associated with siRNA-mediated gene silencing. We obtained deletion alleles
in F37C4.5, hgo-1, and math-33 from the Caenorhabditis Genetics Center (CGC) and the
National Bioresource Project of Japan, and generated new deletion alleles in simr-1,
C33G3.6, and Y57G11C.3 by CRISPR. Strains containing mutations in mut-16, other
known mutator complex proteins such as rde-8 or nyn-1; nyn-2, or the RNAi-related
protein, rsd-2, are defective in both somatic and germline exogenous RNAi (Figure 2A)
(Zhang et al. 2012; Tsai et al. 2015; Sakaguchi et al. 2014; Han et al. 2008; Tijsterman et
al. 2004). To determine whether any of the MUT-16-associated proteins play a role in
exogenous RNAi, we tested the deletion alleles on both somatic and germline RNAi. All
deletions, including simr-1, elicited RNAi phenotypes similar to wild-type animals
indicating that these genes are not required for exogenous RNAi (Figure 2A). We
hypothesized that simr-1 could be redundant with its paralog, hpo-40, so we additionally
tested hpo-40 single mutants and simr-1; hpo-40 double mutants. Both the single and
double mutants elicited RNAi phenotypes similar to wild-type animals indicating that
neither hpo-40 alone nor the two proteins acting together are required for exogenous
RNAi (Figure 2B).
To assess the levels of endogenous siRNAs in each deletion mutant, we isolated RNA
from synchronous 1-day adult animals and generated small RNA sequencing libraries.
Because these proteins were identified by MUT-16 IP-mass spec, we focused on a group
of approximately 2000 genes that are known targets of the mutator pathway (Lee et al.
25
2012; Gu et al. 2009; Phillips et al. 2014; Zhang et al. 2011; Tsai et al. 2015). We observed
a substantial reduction in total small RNAs mapping to these mutator-target genes when
known components of the mutator complex or RNA silencing pathway, such as mut-16,
wago-1, rde-8, or nyn-1; nyn-2 are disrupted (Figure 2C). We also observed a reduction
in small RNAs mapping to the mutator-target genes, albeit more modest, in math-33 and
simr-1 mutants (Figure 2C-D). However, due to asynchrony and slow growth of the math-
33 mutant animals that could confound the data analysis, we chose not to further analyze
the libraries made from this strain at this time. In contrast to the mutator-target genes, we
observed no change in total small RNAs mapping to CSR-1-target genes in the simr-1
mutant (Figure 2E). To test for redundancy between simr-1 and its paralog, hpo-40, in the
endogenous siRNA pathway, we additionally examined levels of small RNAs mapping to
mutator-target genes in hpo-40 single mutants and simr-1; hpo-40 double mutants. We
observed no significant reduction in mutator-dependent small RNAs in the hpo-40 single
mutant, and the reduction in mutator-dependent small RNAs in the simr-1; hpo-40 double
mutant resembled that of the simr-1 single mutant (Figure 2-figure supplement 1A-B).
Therefore, we concluded that SIMR-1 alone is required for siRNA production at some
mutator-target genes.

simr-1 mutants have a mortal germline at elevated temperature
Mutations in the mutator pathway are temperature-sensitive sterile, while mutations in
other related small RNA pathways have a variety of fertility defects (Ketting et al. 1999;
Zhang et al. 2011). For example, mutations in the C. elegans ortholog of Piwi, prg-1,
which associates with piRNAs, display a progressive sterility that accumulates over many
26
generations (also referred to as a Mortal Germline or Mrt phenotype), and mutations in
nuclear RNAi pathway genes nrde-1, nrde-2, nrde-4, and hrde-1 or in the rsd-2 and rsd-
6 genes elicit a similar Mrt phenotype, but only at elevated temperature (Simon et al.
2014; Sakaguchi et al. 2014; Buckley et al. 2012). In order to determine if simr-1 mutants
have fertility defects or the Mrt phenotype observed in many other small RNA silencing
pathway mutants, we quantified their brood size at 20°C, and after every generation at
25°C for 11 generations. mut-16 mutants were included as a control and, as expected,
fertility was reduced by 95.3% in the first generation at 25°C, with the few fertile animals
producing only sterile progeny by the second generation at 25°C (Figure 3A). In contrast,
wild-type animals displayed a 40.3% reduction in brood size and simr-1 mutants displayed
a 59.0% reduction in brood size after a single generation at 25°C compared to 20°C
(Figure 3A). However, unlike wild-type animals which remained fertile after more than 11
generations at 25°C, simr-1 mutants became progressively sterile over the next 10
generations at 25°C until reaching complete sterility at generation 11 (Figure 3A). We
additionally tested the fertility of the hpo-40 single mutant, which was indistinguishable
from wild-type, and the simr-1; hpo-40 double mutant which became sterile after
approximately 11 generations, similar to the simr-1 single mutant (Figure 3-figure
supplement 1). These data indicate that loss of simr-1 at elevated temperature triggers a
molecular defect that is cumulative and ultimately results in loss of fertility.
Because small RNA pathways play key roles in the regulation of transposons, one
hypothesis would be that increased DNA mutations triggered by transposon mobilization
in simr-1 mutants at 25°C lead to reduced fertility over the course of multiple generations.
To address this possibility, we selected wild-type and simr-1 mutant animals raised for 10
27
generations at 25°C, and returned them to 20°C. Within approximately four generations
at 20°C, the fertility of simr-1 mutants recovered to within 72.8% of pre-25°C levels (Figure
3B). These data indicate that the reduction in simr-1 fertility at 25°C is not primarily due
to the accumulation of DNA mutations, but may be due to transcriptional or chromatin
changes that can be reset after recovery at 20°C, similar to what has been observed
previously for hrde-1 and hrde-2 (Spracklin et al. 2017; Ni et al. 2016).

simr-1 Mrt phenotype results from defective sperm and oocytes
To determine whether the Mrt phenotype observed in simr-1 mutants at 25°C is due
to defects in oogenesis or spermatogenesis we conducted mating assays. First, we
crossed wild-type or simr-1 mutant males raised at 20°C, a single generation at 25°C, or
after 10 generations at 25°C to fog-2 females, which cannot make their own sperm, raised
at 20°C. simr-1 mutant males raised for a single generation at 25°C sired fewer progeny
than the wild-type control males, and simr-1 mutant males raised for 10 generations at
25°C were nearly sterile, similar to simr-1 hermaphrodites raised for 10 generations at
25°C (Figure 3C). We next sought to address whether simr-1 mutant oocytes are similarly
compromised. Males expressing fluorescently tagged pgl-1::gfp (Andralojc et al. 2017),
were mated to simr-1 mutant hermaphrodites raised at 20°C, a single generation at 25°C,
or after 10 generations at 25°C. The pgl-1::gfp males were used to easily distinguish
between cross progeny and self progeny from the simr-1 mutant hermaphrodites. simr-1
mutant hermaphrodites raised for a single generation at 25°C and provided with wild-type
sperm produced a similar number of progeny to a wild-type control. In contrast, after 10
generations at 25°C, simr-1 mutant hermaphrodites were nearly sterile, even when
28
provided with wild-type sperm (Figure 3D). These data indicate that both
spermatogenesis and oogenesis are defective in simr-1 mutants raised at elevated
temperature for multiple generations.

simr-1 Mrt phenotype is associated with increased levels of germ cell apoptosis
Apoptosis occurs in the late pachytene region of the germline where approximately
half of all germ cells are eliminated by physiological apoptosis in a wild-type animal
(Gumienny et al. 1999). DNA damage or other stressful conditions can trigger an
increase in apoptosis as part of a quality control mechanism (Gartner et al. 2000; 2008).
To determine if simr-1 mutant gonads have increased apoptosis, we introduced the
CED-1::GFP reporter, which allows visualization of apoptotic germ cells, into the simr-1
mutant (Schumacher et al. 2005). We observed no significant differences in apoptotic
germ cells at 20°C (Figure 3E). After a single generation at 25°C, we observe a
dramatic increase in apoptotic germ cells, with apoptosis levels modestly higher in wild-
type compared to simr-1 mutants. This spike in apoptotic germ cells in the first
generation at 25°C is followed by a reduction in apoptosis in the second generation at
25°C. However, only after 10 or 11 generations at 25°C does the number of apoptotic
germ cells in simr-1 mutants rise significantly compared to wild-type animals (Figure
3E). These data suggest that an increase in germ cell dysfunction in simr-1 mutant
animals after multiple generations of growth at 25°C is associated with both increased
germ cell apoptosis and reduced fertility. Nonetheless, it is important to note that similar
levels of apoptotic germ cells are observed in fertile wild-type animals after only one
29
generation at 25°C, indicating that a high level of apoptosis is not always directly
correlative with sterility.

Mutations in simr-1 desilence a piRNA sensor but not an ERGO-1-dependent siRNA
sensor
In a previously described mutagenesis screen, we identified novel genes acting in the
piRNA-mediated silencing pathway using a strain expressing GFP::H2B carrying a piRNA
target in its 3’UTR (the “piRNA sensor”) (Bagijn et al. 2012; de Albuquerque et al. 2014).
Because the piRNA sensor is subject to siRNA-mediated heritable silencing (RNAe)
making it no longer susceptible to desilencing when the piRNA pathway is compromised,
the screen was performed in a henn-1 mutant background, which partially desilences this
transgene and allows for the identification of both piRNA pathway and secondary siRNA
pathway mutants (Kamminga et al. 2012). From this screen we identified two alleles of
simr-1 that further desilence the piRNA sensor transgene in the henn-1 mutant
background (Figure 4A). The first, simr-1[A11V], is found in a well-conserved region near
the N-terminus of the protein and the second, simr-1[R159C], is the absolutely conserved
arginine that plays a structural role in the extended Tudor domain (Figure 1E).
Interestingly, when we crossed our simr-1 deletion mutant into the piRNA sensor strain
without the henn-1 mutant, we observed that simr-1 was not sufficient to desilence the
piRNA sensor transgene in the absence of the henn-1 mutant (Figure 4-figure supplement
1A), similar to what has been observed previously with prg-1 (Luteijn et al. 2012). In
contrast, a mutation in mut-16 robustly desilences the same piRNA sensor transgene
(Figure 4-figure supplement 1A). These data indicate that a mutation in simr-1, like prg-
30
1, is sufficient to desilence a sensitized piRNA sensor strain, but cannot reactivate a
piRNA sensor silenced by RNAe.
To examine the role of SIMR-1 in other small RNA pathways, we next introduced a
simr-1 mutant into the 22G-siR1 sensor which is sensitive to perturbations in the ERGO-
1 26G-siRNA pathway and the downstream mutator pathway (Montgomery et al. 2012).
A mutation in simr-1 was unable to desilence the 22G-siR1 sensor (Figure 4-figure
supplement 1B). In contrast, a mutation in mut-16 robustly desilenced the 22G-siR1
sensor (Figure 4-figure supplement 1B). Furthermore, when animals with mutations in the
ERGO-1 26G-siRNA pathway, like eri-7 (Fischer et al. 2008), are fed lir-1, hmr-1, or dpy-
13 double-strand RNA, they display an Enhanced RNAi (Eri) phenotype which was not
observed with the simr-1 mutant (Figure 4-figure supplement 1C). These data indicate
that SIMR-1 is not required for silencing of genes targeted by the ERGO-1 26G-siRNA
pathway.

SIMR-1 is required to prevent sterility after reestablishing WAGO-class 22G-siRNA
production
Neither the mutator pathway nor the piRNA pathway are essential for fertility under
normal growth conditions (Ketting et al. 1999; Zhang et al. 2011; Batista et al. 2008; Wang
and Reinke 2008; Simon et al. 2014). Nonetheless, restoration of the mutator pathway,
and therefore RNA silencing by WAGO-class 22G-siRNAs, in a strain lacking both the
mutator pathway and the piRNA pathway, causes sterility (de Albuquerque et al. 2015;
Phillips et al. 2015). This sterility is a direct result of the routing of essential genes into the
mutator pathway and indicates that inheritance of piRNAs from one generation to the next
31
is critical to ensuring that the correct genes are silenced by the mutator pathway. To
determine whether simr-1, like prg-1, is required to maintain fertility when resetting the
mutator pathway, we crossed two strains to one another containing distinct mutations in
the mutator pathway, mut-7 and mut-14 smut-1, such that their progeny would inherit a
wild-type copy of mut-7 from one parent, a wild-type copy mut-14 smut-1 from the other,
and thus would be competent to produce WAGO-class 22G-siRNAs (Figure 4B). The
hermaphrodite strain always additionally carried the unc-119 mutation, which allowed us
to easily distinguish between self progeny which have the Uncoordinated (Unc)
phenotype and cross progeny which have wild-type movement. If simr-1 is required for
the proper functioning of the piRNA pathway, we would predict that when it, like prg-1, is
introduced into the two strains used to reset the mutator pathway the progeny of the cross
will be sterile. In fact, this result is what we observed. In the control cross (mut-14 smut-
1 males mated to mut-7 unc-119 hermaphrodites), only 13.0% of the F1 heterozygous
progeny were sterile (Figure 4B). In contrast, when the simr-1 mutation is present in both
parental strains (simr-1; mut-14 smut-1 males mated to simr-1; mut-7 unc-119
hermaphrodites) the percentage of sterile progeny increased to 47.1%, and for the prg-1
cross (prg-1; mut-14 smut-1 males mated to prg-1; mut-7 unc-119 hermaphrodites), the
number of sterile animals increases further to 98.8% (Figure 4B). These results indicate
that simr-1, like prg-1, is required during establishment of the mutator pathway to promote
fertility, likely by directing mutator-dependent silencing to piRNA-targeted genes.

32
The Tudor domain of SIMR-1 is required for its localization and function
To determine whether the Tudor domain of SIMR-1 is necessary for its localization to
germline foci, we used CRISPR to engineer the R159C mutation into the simr-1::gfp
strain. The R159C allele, isolated from a mutagenesis of the henn-1; piRNA sensor strain,
is predicted to disrupt the conformation of the extended Tudor domain (Liu et al. 2010a).
By live imaging, we observed that SIMR-1[R159C]::GFP no longer forms germline foci,
despite its clear expression in the cytoplasm of germ cells (Figure 4C). We further
confirmed that SIMR-1[R159C]::GFP is expressed at wild-type levels by western blot
(Figure 4-figure supplement 1D). These data indicate that an intact extended Tudor
domain is not required for SIMR-1 expression but is essential for the localization of SIMR-
1 to germline foci.
We next investigated whether the simr-1[R159C]::gfp strain exhibited fertility defects
at elevated temperature. Like the simr-1 deletion allele, simr-1[R159C]::gfp exhibited
progressive sterility at elevated temperature, becoming sterile after approximately 10-11
generations (Figure 4D). In contrast, the wild-type simr-1::gfp remained fertile for the
duration of the experiment (Figure 4D). Together, these data show that the extended
Tudor domain is essential for SIMR-1 function, and that disruption of the Tudor domain
results in loss of SIMR-1 germline foci and causes a Mrt phenotype similar to that of the
simr-1 deletion allele.

SIMR-1 is required for small RNA production at piRNA-target genes
To comprehensively characterize the role of SIMR-1 in C. elegans endogenous small
RNA pathways, we generated small RNA libraries from wild-type and simr-1 mutants at
33
20°C and after culturing for one, two, seven, or 10 generations at 25°C. For comparison,
we also generated small RNA libraries from wild-type, mut-16, and prg-1 mutants at 20°C
and from wild-type and mut-16 mutants cultured for a single generation at 25°C. In simr-
1 mutants, 817 genes were depleted of small RNAs and 213 genes were enriched for
small RNAs at 20°C when compared to wild-type at 20°C (Figure 5A and Supplementary
File 3, Manage et al 2020). After one generation at 25°C, 1258 genes were depleted of
small RNAs and 2712 genes were enriched for small RNAs compared to wild-type also
cultured for one generation at 25°C (Figure 5A and Supplementary File 3, Manage et al
2020). When simr-1 mutants were then cultured for two, seven, or 10 generations at 25°C,
927, 885, and 907 genes were depleted of small RNAs and 194, 110, and 100 genes
were enriched for small RNAs, respectively, when compared to both wild-type cultured at
25°C for one generation and wild-type cultured at 25°C in parallel to simr-1 for an equal
number of generations (Figure 5A and Supplementary File 3, Manage et al 2020). These
data implicate SIMR-1 in the production or maintenance of small RNAs at many C.
elegans genes.
siRNAs can be classified based on their Argonaute protein binding partner and the
other proteins or protein complexes required for their biogenesis. To identify the small
RNA pathway(s) in which SIMR-1 plays a role, we looked at the change in total small
RNA levels at groups of genes known to be targets of the CSR-1, mutator, piRNA, or
ERGO-1 pathways in simr-1 mutants compared to wild-type at both 20°C and a single
generation at 25°C (Lee et al. 2012; Fischer et al. 2011; Gu et al. 2009; Phillips et al.
2014; Zhang et al. 2011; Tsai et al. 2015). Small RNAs derived from CSR-1-target genes
were modestly up-regulated at 20°C and more dramatically up-regulated after a single
34
generation 25°C in simr-1 mutants (Figure 5B-C and Figure 5-figure supplement 1A). In
contrast, small RNAs from mutator-target genes and piRNA-target genes were reduced
in simr-1 mutants at both 20°C and 25°C (Figure 5B-C and Figure 5-figure supplement
1A). piRNA target genes make up the majority of mutator-target genes (Figure 5D). To
determine if piRNA-target genes are more severely reduced of small RNAs in simr-1
mutants than other mutator-target genes, we generated a list of mutator-target genes
whose small RNAs are either unchanged or increased in prg-1 mutants (log2(fold change
small RNA abundance) ≥ 0 in prg-1 mutants relative to wild-type). These PRG-1-
independent mutator-target genes are not reduced of small RNAs compared to all siRNA
target genes and are significantly less depleted of small RNAs compared to all mutator-
target genes or piRNA-target genes (Figure 5B). Furthermore, the well-characterized
endogenous RDE-1 target, Y47H10A.5 (Corrêa et al. 2010), was not depleted of small
RNAs in simr-1 mutants at either 20°C or 25°C or in prg-1 mutants at 20°C, but was
severely depleted of small RNAs in mut-16 mutants at both 20°C and 25°C (Figure 5-
figure supplement 1B), demonstrating that like exogenous RNAi targets (Figure 2A), small
RNA levels at endogenous RDE-1 targets are not affected in the simr-1 mutant. Small
RNAs from ERGO-1 target genes were reduced mildly at 20°C and more severely at 25°C
(Figure 5B and Figure 5-figure supplement 1A), however because simr-1 was unable to
desilence the 22G-siRNA sensor and did not have an Eri phenotype (Figure 4-figure
supplement 1B-C), we did not pursue further investigation of the ERGO-1 pathway.
Therefore, these data indicate that SIMR-1 is important for the production of high levels
of endogenous small RNAs at many mutator-target genes, including primarily piRNA-
35
target genes, but is not required for small RNA production at CSR-1-target genes or at
endogenous and exogenous RDE-1-target genes.

SIMR-1 is not required for piRNA biogenesis or stability
84% of genes with reduced small RNAs in a simr-1 mutant at 20°C also have reduced
small RNAs in a prg-1 mutant at 20°C (Figure 5D). This reduction of siRNAs at piRNA-
target genes could result from a loss of piRNAs in the simr-1 mutant animals, or
alternatively, piRNAs could be expressed at wild-type levels and only the downstream
siRNAs could be affected. To address these possibilities, we counted the number of reads
mapping to annotated piRNA loci in wild-type and simr-1 mutants. Similarly to what has
been previously reported, piRNA expression is significantly reduced at 25°C compared to
20°C in wild-type animals (Bélicard et al. 2018). However, we observed no significant
difference between total piRNA levels in simr-1 mutants compared to wild-type animals
at either temperature (Figure 5E). We next determined whether individual piRNAs are
increased or reduced in expression in simr-1 mutants. In contrast to prg-1 mutants in
which 83% of piRNAs are reduced by at least two-fold, in simr-1 mutants less than 1% of
piRNAs are reduced by at least two-fold (Figure 5F). We next identified predicted piRNA
target genes for the piRNAs that were reduced by at least two-fold in simr-1 mutants
(Shen et al. 2018; Zhang et al. 2018a; Wu et al. 2018; 2019). Specifically, we selected
genes predicted to be targets for our simr-1-depleted piRNAs by piRTarBase using
relaxed piRNA targeting rules and identified by CLASH data (Supplementary File 4,
Manage et al 2020). Of the 37 predicted target genes for our simr-1-depleted piRNAs,
only five have reduced small RNAs in simr-1 mutants (Supplementary File 4, Manage et
36
al 2020), indicating that the simr-1-depleted piRNAs are not a major driver of siRNA
depletion in simr-1 mutants. These data together indicate that SIMR-1 functions
downstream of piRNA biogenesis.

Small RNAs are progressively depleted across generations from some piRNA-
target loci at 25°C
Because simr-1 mutant animals become sterile after approximately 10 generations at
25°C, we next examined how the levels of small RNAs generated from mutator and
piRNA-target genes change after two, seven, or 10 generations at 25°C, compared to a
single generation at 25°C. At each generation, we compared the genes that lose small
RNAs by at least two-fold in the simr-1 mutant to genes that lose small RNAs by at least
two-fold in mut-16 mutants at 25°C, and to prg-1 mutants at 20°C. At all generations,
SIMR-1-dependent siRNA target genes largely overlapped with mut-16-dependent siRNA
target genes. Specifically, 80%, 88%, 97% and 89% of SIMR-1-dependent small RNA
target genes at 25°C for one, two, seven, and 10 generations are reduced of small RNAs
in mut-16 mutants, respectively, compared to 84% for SIMR-1-dependent siRNA target
genes at 20°C (Figure 5D and Figure 5-figure supplement 1C). We next examined the
overlap of SIMR-1-dependent small RNA target genes with prg-1 mutants at 20°C. 55%,
64%, 72%, 70% of SIMR-1-dependent small RNA target genes at 25°C for one, two,
seven, and 10 generations are reduced of small RNAs in prg-1 mutants at 20°C,
respectively, compared to 84% for SIMR-1-dependent siRNA target genes at 20°C
(Figure 5D and Figure 5-figure supplement 1C). While the overlap of SIMR-1-dependent
small RNA target genes with piRNA-dependent small RNA target genes is reduced at
37
25°C compared to 20°C, at least some of this difference may be attributed to the
sequencing of prg-1 mutant small RNA libraries from animals raised at 20°C only. In fact,
the total number of genes reduced of small RNAs in simr-1 mutants that overlap with
piRNA-target genes remains similar between temperatures and across generations
(Figure 5D and Figure 5-figure supplement 1C). However, while the number of piRNA-
target genes that lose small RNAs in a simr-1 mutant doesn’t change significantly with
temperature or later generations, we do observe a modest but significant progressive
reduction in the number of small RNAs mapping to all piRNA-target genes corresponding
to the number of generations at 25°C (Figure 5-figure supplement 1D). Because the
number of simr-1-target genes does not become substantially greater after 10
generations at elevated temperature, these data indicate that the observed sterility is not
due to loss of small RNAs from more loci after 10 generations. Furthermore, while many
piRNA-target genes become more depleted of small RNAs after 10 generations at
elevated temperature, this loss of small RNAs is unlikely to be a contributing factor to the
progressive loss of fertility in these animals because small RNA loss is even more severe
in fertile prg-1 mutants at 20°C (Fig 5C and S5D).

SIMR-1 is required for small RNA production at many piRNA-targeted transposons
and repetitive elements
The mutator pathway is required for the production of siRNAs at many transposons
and repeat loci, and in the absence of mut-16 or other mutator complex proteins
transposon activity has been detected for at least seven distinct families of DNA
transposons (Tc1-Tc5, Tc7, CemaT1) (Eide and Anderson 1985; Collins et al. 1989; Levitt
38
and Emmons 1989; Yuan et al. 1991; Collins and Anderson 1994; Rezsohazy et al. 1997;
Bessereau 2006; Brownlie and Whyard 2004). In contrast, only a single transposon
family, Tc3, has been demonstrated to transpose upon loss of the piRNA machinery,
though several other DNA transposon loci are up-regulated at the mRNA level or lose
mutator-dependent siRNAs (Das et al. 2008; Bagijn et al. 2012; McMurchy et al. 2017;
Wallis et al. 2019; Reed et al. 2020). To address the role of SIMR-1 in the regulation of
transposons and repeat loci, we first defined a list of mut-16-dependent transposons and
repeats using a cutoff of two-fold reduction of small RNAs in the mut-16 mutant compared
to wild-type at 20°C. All features also met the requirements of having at least 10 RPM in
either mutant or wild-type and a DESeq2 adjusted p-value of ≤ 0.05. Of these mut-16-
dependent transposons and repeats, 11% and 25% of transposons at 20°C and 25°C
respectively, and 35% and 45% of repeat loci, at 20°C and 25°C respectively, were
reduced by two-fold or greater of small RNAs in simr-1 mutants compared to wild-type
(Figure 6-figure supplement 1A). Furthermore, 82% of the mut-16-dependent
transposons depleted of small RNAs by greater than two-fold in simr-1 mutants at 20°C
were also depleted in prg-1 mutants at 20°C (Figure 6-figure supplement 1B). Similarly,
80% of the mut-16-dependent repeats depleted of small RNAs by greater than two-fold
in simr-1 mutants at 20°C were also depleted in prg-1 mutants at 20°C (Figure 6-figure
supplement 1B). We next focused on transposons for which silencing is known to be
either piRNA-dependent or piRNA-independent. Transposon Tc3 becomes active in
mutants of the mutator pathway and the piRNA pathway, while Tc1 and Tc4 activity is
specific to the mutator pathway (Das et al. 2008). Tc2 activity has not been measured in
piRNA pathway mutants, but the Tc2 transposase mRNA is significantly up-regulated in
39
a prg-1 mutant (Wallis et al. 2019). We next determined the number of small RNAs
mapping to these four transposon sequences in simr-1 mutants compared to wild-type.
Small RNAs mapping to Tc2 and Tc3 were significantly reduced in both the simr-1 mutant
as well as in a mut-16 mutant, at both 20°C and 25°C (Figure 6A and Figure 6-figure
supplement 1C). In contrast, small RNAs mapping to Tc1 and Tc4v, the variant of Tc4
containing the Tc4 transposase mRNA sequence (Li and Shaw 1993), were not reduced
in simr-1 mutants (Figure 6A and Figure 6-figure supplement 1D). These data indicate
that SIMR-1 is required for small RNA production or maintenance at piRNA-targeted
transposons but not at transposons targeted independently of piRNAs.

simr-1 mutants have increased levels of small RNAs mapping to histone genes
We next focused on the genes for which the mapped small RNAs increase in simr-1,
prg-1 and mut-16 mutants. In general, fewer genes have a two-fold increase in small
RNAs compared to a two-fold decrease in small RNAs for simr-1, prg-1 and mut-16
mutants at 20°C (Figure 5A). These data would indicate that the SIMR-1, along with PRG-
1 and MUT-16, plays a more significant role in production or maintenance of small RNAs
rather than in suppression of small RNA production. Nonetheless, 213 genes gain small
RNAs by greater than two-fold in simr-1 mutants, 49% of which also gain small RNAs in
prg-1 mutants (Figure 5A and 6B). Interestingly only three of these genes (1%) also gain
small RNAs in mut-16 mutants (Figure 6B). While manually examining the list of genes
enriched for small RNAs in both simr-1 and prg-1 mutants at 20°C, we noticed that this
list included numerous histone genes. Of the 104 genes enriched for small RNAs in both
simr-1 and prg-1 mutants, 28 are histone genes (Figure 6C) (Pettitt et al. 2002). These
40
28 genes make up 38% of all C. elegans histone genes (Figure 6C). An additional 30
histone genes (41%) are enriched for small RNAs in only prg-1 mutants, and only one
histone gene is enriched for small RNAs in both mut-16 and prg-1 mutants (Figure 6C).
Overall, histone genes are highly enriched for small RNAs in both simr-1 and prg-1
mutants, though this enrichment is lessened across multiple generations at 25°C,
suggesting that it may not be associated with the sterility phenotype (Figure 6D).
Nonetheless, this enrichment of small RNAs at histone genes in both simr-1 and prg-1
mutants is clearly in contrast to mut-16 mutants at 20°C and 25°C, where the majority of
histone genes are unchanged or depleted of small RNAs (Figure 6D). We further
examined the histone genes by histone gene class and we observed that some histone
genes classes such as H2A and H3 genes are enriched for small RNAs in both simr-1
and prg-1, whereas others such as H2B are enriched for small RNAs primarily in prg-1
mutants (Figure 6E and Figure 6-figure supplement 1E). This increase in small RNA
production to histone genes has been observed previously in prg-1 mutants and these
histone-derived small RNAs are dependent on the mutator complex for their biogenesis
(Barucci et al. 2020; Reed et al. 2020). These data suggest that enrichment of small RNAs
at certain classes of histone genes is a signature unique to the simr-1 and prg-1 mutants
and not the mutator pathway, and thus provides additional evidence that SIMR-1 plays a
key role in the piRNA pathway.

Most SIMR-1-target genes are not desilenced in a simr-1 mutant
To determine whether the observed changes to small RNA levels alter gene
expression in simr-1 mutants, we next sequenced mRNAs isolated from wild-type, simr-
41
1 mutant and mut-16 mutant animals at 20°C and from wild-type and simr-1 mutant
animals after one, two, seven, or 10 generations at 25°C. We identified 139 genes whose
mRNA expression was reduced by at least two-fold in simr-1 mutants at 20°C and 164
genes whose mRNA expression was increased by at least two-fold in simr-1 mutants at
20°C (Figure 6F and Supplementary File 5, Manage et al 2020). Not surprisingly, the simr-
1 up-regulated genes were enriched for mutator-target genes and PRG-1-target genes,
which initially suggested to us that there may be a direct correlation between loss of small
RNAs and an increase in mRNA expression at some loci (Figure 6G). However, when we
directly compared the list of genes with increased mRNA expression in a simr-1 mutant
(164 genes) to the genes with reduced small RNAs in a simr-1 mutant (817 genes) we
found only 18 genes in common and, furthermore, we do not see a significant change in
mRNA expression for the genes depleted of small RNAs in simr-1 mutants (Figure 6H
and Figure 6-figure supplement 1F). Similarly, in mut-16 mutants, we do not observe a
substantial change in mRNA expression for the genes depleted of small RNAs (Figure
6H), which is consistent with recent findings that the majority of mutator-target genes and
PRG-1-target genes are not desilenced in mut-16 or prg-1 mutants, respectively (Barucci
et al. 2020; Reed et al. 2020). We also observed a modest enrichment of spermatogenic
genes among the simr-1 up-regulated genes. This result is similar to the previously
published observation that spermatogenesis genes are upregulated in prg-1 and mut-16
mutants (Reed et al. 2020; Rogers and Phillips 2020), and is consistent with simr-1 acting
with prg-1 in the regulation of PRG-1 target genes. These data indicate that the majority
of SIMR-1-target genes are not derepressed in a simr-1 mutant, which suggests that
either SIMR-1-dependent siRNAs are required only to initiate but not maintain silencing
42
of their targets or that additional layers of regulation maintain silencing of these genes in
the absence of SIMR-1-dependent siRNAs.
We next focused on the genes down-regulated in simr-1 mutants and found that those
genes were also enriched for mutator-target genes and PRG-1-target genes (Figure 6G),
indicating that some mutator and PRG-1-target genes are up-regulated, while others are
down-regulated in simr-1 mutants. When we looked exclusively at the genes enriched for
small RNAs in simr-1 mutants, we observed a modest down-regulation of these genes at
the mRNA level (Figure 6H), indicating that the small RNA gained in the simr-1 mutant
are sufficient to promote down-regulation of their target mRNAs. The same trend was not
observed for genes enriched for small RNAs in mut-16 mutants (Figure 6H). Histone
genes, including H2A and H3, were amongst those genes enriched for small RNAs and
with reduced mRNA expression in simr-1 mutants (Figure 6D-E, 6I, and Figure 6-figure
supplement 1G), similar to what has previously been observed in prg-1 mutants (Barucci
et al. 2020; Reed et al. 2020). We hypothesize that the small RNAs gained in simr-1
mutants may depend on the mutator pathway, similar to what has been shown for the
small RNAs targeting histone genes in the prg-1 mutant (Barucci et al. 2020; Reed et al.
2020), and therefore these small RNAs are competent to silence their target mRNAs. In
contrast, the mutator pathway is non-functional in the mut-16 mutant, therefore the small
RNAs gained in this mutant are likely to be a distinct class of small RNAs, possibly CSR-
1-class siRNAs, which do not generally silence their mRNA targets (Claycomb et al. 2009;
Wedeles et al. 2013).
Finally, to determine whether the sterility observed in simr-1 mutants raised at 25°C
for 10 generations could be attributed to gene expression changes, we looked for mRNAs
43
up or down-regulated in simr-1 mutants raised at 25°C for 10 generations compared to
wild-type raised under the same conditions that were not up or down-regulated in simr-1
mutants raised at 20°C or in simr-1 mutants raised at 25°C for only a single generation
(exclusive to gen. 10). We identified only 34 genes significantly down-regulated
exclusively at generation 10 and 112 genes significantly up-regulated exclusively at
generation 10 (Supplementary File 5, Manage et al 2020). The genes up-regulated
exclusively in simr-1 mutants after 10 generations were enriched for mutator-target
genes, PRG-1-target genes, ALG-3/4-target genes and spermatogenic genes while the
down-regulated genes were not enriched for any gene list that we examined (Figure 6G).
While these up-regulated genes are exclusive to 10 generations at 25°C, the classes of
enriched genes (mutator targets, PRG-1 targets, and spermatogenic genes) are similar
to what was observed in simr-1 mutants at 20°C. While we cannot attribute the sterility
observed in these animals directly to the misregulation of any specific genes, we
hypothesize that an increase in the expression of spermatogenesis genes during
oogenesis, along with the expression of mutator and PRG-1-target genes could contribute
to germ cell dysfunction.

SIMR-1 forms foci near Mutator foci, P granules and Z granules
P granules, Mutator foci, and Z granules are all phase-separated biomolecular
condensates which lie adjacent to one another at the nuclear periphery (Uebel et al. 2018;
Wan et al. 2018; Brangwynne et al. 2009). From live imaging of fluorescently-tagged
SIMR-1 and MUT-16, we observed that SIMR-1 forms foci closely associated with Mutator
foci (Figure 1C), however from this preliminary analysis we were unable to conclude
44
whether they fully colocalized. To first address the spatial relationship between SIMR-1
and MUT-16, we immunostained fluorescently-tagged SIMR-1 and MUT-16. We
observed that SIMR-1 foci are closely associated with Mutator foci (96.4% of the time with
no empty space between fluorescent signals, n=56 SIMR-1 foci), however they do not
fully colocalize suggesting that they are distinct structures (Figure 7A). This result is
supported by our previous observation that SIMR-1 foci are not disrupted in a mut-16
mutant, nor are Mutator foci disrupted by the simr-1; hpo-40 double mutant (Figure 1D).
Furthermore, we have not been able to unambiguously co-immunoprecipitate MUT-16
with SIMR-1, which indicates that, despite our initial identification of SIMR-1 in the MUT-
16 IP-mass spectrometry experiment, the physical interaction between these two proteins
may be weak or transient.
Both P granules and Z granules are closely associated with Mutator foci (Wan et al.
2018; Phillips et al. 2012), so we next asked whether SIMR-1 foci colocalize with either
PGL-1, marking P granules, or ZNFX-1, marking Z granules. SIMR-1 foci are closely
associated with both P granules and with Z granules (100% of the time with P granules,
n=56 SIMR-1 foci, and 100% of the time with Z granules, n=62 SIMR-1 foci). However,
we found that SIMR-1 foci do not fully colocalize with either structure, and in some cases
multiple SIMR-1 foci can associate with a single focus of another granule type (Figure 7A,
see inset for SIMR-1 and ZNFX-1 localization). SIMR-1 foci do appear to be more closely
associated with Z granules than with P granules and quantification of distances between
fluorescence centers of each foci supports this observation (Figure 7A-B). Because
SIMR-1 promotes siRNA biogenesis at piRNA target genes, we also examined the
colocalization of SIMR-1 and PRG-1, which has previously been shown to localize to P
45
granules (Batista et al. 2008; Wang and Reinke 2008). Similar to what we observed with
PGL-1, PRG-1 is localized adjacent to but not coincident with SIMR-1 foci (Figure 7-figure
supplement 1A). Furthermore, SIMR-1 is not required for PRG-1 localization or
expression, and PRG-1 is not required for SIMR-1 localization (Figure 7-figure
supplement 1B-C). These data indicate that, while SIMR-1 and PRG-1 function in the
same pathway to mediate siRNA biogenesis at piRNA target genes, they do not colocalize
and are not required for one another’s localization or expression.
Also identified in our MUT-16 and SIMR-1 immunoprecipitations was RSD-2, a
previously characterized RNAi factor required for exogenous RNAi introduced in low
doses and production of secondary siRNAs at target genes dependent on the ERGO-1
primary siRNA pathway. Because RSD-2 also forms foci in close proximity to Mutator foci
(Figure 1-figure supplement 1C), we next generated a strain with fluorescently-tagged
SIMR-1 and RSD-2. Following immunostaining, we observed that SIMR-1 and RSD-2
were highly coincident, suggesting SIMR-1 and RSD-2 may localize to the same
perinuclear structure (Figure 7A-B). These results indicate that SIMR-1 and RSD-2
interact closely with one another at perinuclear foci near but distinct from Mutator foci, P
granules and Z granules. Because these foci are distinct from previously characterized
structures, we are calling them SIMR foci.
Finally, to better understand the organization of these multiple perinuclear foci, we
immunostained for SIMR-1, ZNFX-1, and PGL-1 together. Interestingly, we observed that
the foci appeared to be stacked, with ZNFX-1 localizing between SIMR-1 and PGL-1
(Figure 7C). This result is reminiscent of the tripartite PZM granule (P granule/Z
granule/Mutator foci) observed by Wan et al (2018), except that we observe the Z granule
46
flanked by SIMR foci and P granules, instead of Mutator foci and P granules. Therefore,
our data suggest that there are at least four separate compartments at the nuclear
periphery in C. elegans germ cells, that together constitute C. elegans nuage, each with
unique protein components and a distinct molecular role in the RNA silencing pathway.

Discussion
C. elegans utilize the highly abundant siRNAs synthesized by the mutator complex to
reinforce silencing initiated by the piRNA pathway. Here we identify a Tudor domain
protein, SIMR-1, required to mediate effective production of siRNAs from many piRNA-
target mRNAs. We demonstrate that SIMR-1 has a phenotype similar to that of PRG-1,
in that simr-1 mutants can desilence a sensitized piRNA sensor and SIMR-1 is required
to prevent sterility after reestablishing WAGO-class 22G-siRNA production. However, the
phenotypes associated with simr-1 are often weaker than those of prg-1 (see Figs 4B,
5C-D, 6B-E, S5C-D, and S6B), suggesting that simr-1 is not absolutely required to
mediate siRNA amplification at all piRNA target genes, or it acts cooperatively with other
pathways or proteins. SIMR-1 is not RNAi-defective, it cannot desilence a piRNA sensor
silenced by RNAe, and it cannot desilence the ERGO-1-dependent siRNA sensor, all
phenotypes associated with the downstream mutator pathway. Furthermore, siRNAs are
reduced at many piRNA-target loci in simr-1 mutants, but piRNAs themselves are
unaffected. Like PRG-1 and the mutator complex, SIMR-1 forms foci near the nuclear
periphery of germ cells, and while these perinuclear condensates are adjacent to one
another, they all appear to be distinct substructures. Thus, our work identifies a novel
player acting at a step in between piRNA biogenesis and siRNA amplification by mutator
47
complex and suggests a role for multiple perinuclear condensates to promote piRNA-
mediated siRNA production.

Tudor domain proteins in piRNA-mediated silencing
Tudor domain proteins are thought to act as scaffolds in the piRNA pathway, to
engage and assemble multiple partner proteins (Pek et al. 2012). Through promotion of
protein-protein interactions, they can drive piRNA biogenesis and piRNA target silencing.
For example, the Drosophila Tudor domain protein, Krimper, interacts with two Piwi
proteins, Aubergine and Ago3, to coordinate assembly of the ping-pong processing
complex (Webster et al. 2015). Of note, sDMA of Aubergine is required for interaction
with Krimper, but Ago3 can interact with Krimper independently of sDMA, emphasizing
that Tudor domain proteins can play critical roles in the piRNA pathway independent of
sDMA. In fact, like SIMR-1, many human and Drosophila Tudor domain proteins carry
mutations in aromatic cage residues, indicating they may have lost the ability to bind
methylated arginine substrates (Handler et al. 2011; Zhang et al. 2017). For example,
mammalian Tudor domain protein, TDRD2, which is missing one of the four aromatic
cage residues, preferentially recognizes an unmethylated peptide of PIWIL1 over a
dimethylated peptide. This recognition occurs through a negatively charged groove that
occurs at the interface of the canonical Tudor domain and the flanking conserved
elements making up the extended Tudor domain (Zhang et al. 2017). This data would
suggest that Tudor domain proteins that are missing the aromatic cage residues, like
SIMR-1, may still make functional interactions with Piwi proteins or other small RNA
pathway proteins. They may mediate interactions preferentially with unmethylated
48
substrates and/or compete with other Tudor domain proteins for substrates dependent on
methylation status. While we have not yet identified the relevant protein-binding partners
of the SIMR-1 Tudor domain, we hypothesize that they are likely members of the piRNA
pathway or mutator complex and contain the RG repeat motif preferentially recognized
by the Tudor domain. An obvious candidate is PRG-1 itself, as it contains a “GRGRGRG”
sequence near its N-terminus, however there are certainly other candidates and further
experiments will be necessary to test this possibility.

Regulation of piRNA-target genes by perinuclear condensates
While we do not have direct evidence for a physical interaction between SIMR-1 and
PRG-1, it is likely that SIMR-1 interacts with either PRG-1 or some other member of the
piRNA pathway to promote the downstream regulation of piRNA target mRNAs by the
mutator complex. Similarly, because SIMR-1 was initially identified in a MUT-16
immunoprecipitation, it may also interact directly with the mutator complex, even if
transiently. It is therefore interesting that PRG-1, SIMR-1, and the mutator complex all
localize to distinct sub-compartments of nuage (Supplementary File 1, Manage et al
2020). We have observed that Z granules localize between SIMR-1 foci and P granules,
similar to the organization of Mutator foci, Z granules and P granules (Wan et al. 2018).
We have not been able to image SIMR foci with Mutator foci, Z granules, and P granules
simultaneously, so it remains to be determined how these four substructures assemble
together and whether SIMR foci bridge Mutator foci and Z granules, Mutator foci bridge
SIMR foci and Z granules, or whether all three interact. Mutator foci, P granules, and Z
granules all assemble through intracellular phase separation, which brings about the
49
question as to whether SIMR foci may also behave in a liquid-like manner. While we have
not tested this formally, the localization of SIMR-1 nestled among these three other
biomolecular condensates is certainly suggestive. The dynamic nature of these various
condensates could facilitate exchange of RNAs or protein components between
compartments, which may explain how piRNA pathway proteins, SIMR-1, and the mutator
complex could occupy distinct substructures while facilitating regulation of the same
mRNA target genes. Perhaps some proteins have properties making them immiscible in
multiple condensates allowing them to promote transfer of RNAs between compartments,
or alternatively, the exchange of RNAs may occur at their interface.

RSD-2 and SIMR-1 promote the interaction between distinct primary and secondary
siRNA pathways
The colocalization of SIMR-1 and RSD-2 is somewhat surprising given that SIMR-1
and RSD-2 act in distinct small RNA pathways. Specifically, SIMR-1 acts downstream of
PRG-1 in the piRNA pathway and has no defects in exogenous RNAi, whereas RSD-2 is
required to mount an efficient response to exogenous RNAi and silence ERGO-1-target
genes, but is not required for the production of secondary siRNAs at piRNA target genes
(Han et al. 2008; Zhang et al. 2012). While many of their targets are distinct, SIMR-1 and
RSD-2 may play similar roles in mediating the interaction between primary and secondary
siRNA pathways and thus their colocalization may be indicative of a subcellular
compartment mediating this transition between primary and secondary small RNA
pathways.
50
Like RSD-2, the Tudor domain protein RSD-6, the Maelstrom domain protein RDE-
10, the RING-type zinc finger protein RDE-11, and the DEAD box ATPase and Vasa
ortholog RDE-12 likely act downstream of primary Argonaute proteins RDE-1 and ERGO-
1 and are required for the accumulation of mutator-dependent secondary siRNAs (Zhang
et al. 2012; Shirayama et al. 2014; Yang et al. 2014; 2012). Interestingly, there is no data
to suggest that any of these proteins act with SIMR-1 downstream of PRG-1, suggesting
that there could be a completely different set of factors that interact with SIMR-1 at piRNA
targets. While no localization has been determined for RDE-10 and RDE-11, RSD-6
localizes to foci near P granules that may be coincident with SIMR-1 foci. RDE-12
localizes to both RSD-6 foci and P granules, suggesting it can traverse the boundary
between perinuclear condensates, and it has been proposed that RDE-12 may shuttle
primary siRNA bound target mRNAs from P granules to RSD-6 foci to initiate mutator-
dependent siRNA synthesis (Yang et al. 2014). While loss of RDE-12 does not affect
siRNAs mapping at piRNA target genes, there are 36 RDE-12 paralogs in C. elegans,
several of which localize at or near P granules, including GLH-1, -2, -3, -4, DDX-19, LAF-
1, MUT-14, and VBH-1 (Supplementary File 1, Manage et al 2020). One of these proteins
could potentially serve a function similar to RDE-12, in the shuttling of piRNA-targeted
mRNAs into SIMR-1 foci and ultimately to Mutator foci.
In conclusion, numerous proteins have been identified in C. elegans that are required
for piRNA transcription, trimming, and modification (Kamminga et al. 2012; Montgomery
et al. 2012; Billi et al. 2012; Tang et al. 2016; Weick et al. 2014; de Albuquerque et al.
2014; Kasper et al. 2014; Cordeiro Rodrigues et al. 2019; Zeng et al. 2019), however,
how mRNAs travel between the piRNA pathway, required for mRNA recognition, to the
51
mutator pathway, necessary for siRNA production has remained a mystery. Here we
demonstrate that the Tudor domain protein, SIMR-1, is required at a step between the
piRNA pathway to the mutator complex. SIMR-1 may function similarly to how Krimper
coordinates Ago3 and Aubergine during ping pong piRNA biogenesis in Drosophila
(Webster et al. 2015), but in this case bridging the gap between the primary and
secondary phases of the C. elegans piRNA silencing pathway. Finally, SIMR-1 localizes
to cytoplasmic foci near P granules, Z granules, and Mutator foci, implicating a series of
distinct perinuclear condensates in the regulation of mRNAs by the piRNA pathway and
mutator complex.

Materials and Methods
Strains
The C. elegans wild-type strain used is N2. Worms were raised at 20°C according to
standard conditions unless otherwise stated (Brenner 1974). Mutants generated by
CRISPR or obtained from the CGC were outcrossed prior to sequencing or other analysis.
All strains used for this project are listed in Supplementary File 6 (key resources table,
Manage et al 2020).

Plasmid and strain construction
All GFP, mKate2, or mCherry tagged strains were generated by CRISPR genome
editing, with tags inserted at the endogenous locus. simr-1::gfp::3xFLAG repair template
was assembled into pDD282 and mKate2::3xMyc::prg-1 repair template was assembled
into pDD287 (Addgene plasmid # 66823 and #70685) according to published protocols
52
(Dickinson et al. 2015). Design of the mCherry::2xHA plasmid was described previously
(Uebel et al. 2018). The mCherry::2xHA region, which include intronic Floxed Cbr-unc-
119(+), was amplified by PCR and assembled by isothermal assembly with ~1kb of
sequence from either side of the stop codon of the gene to be tagged and the XhoI/EagI
digested pBluescript vector (Gibson et al. 2009). A similar method was used to generate
CRISPR-mediated deletions. A region containing the Floxed Cbr-unc-119(+) was
amplified from the mCherry::2xHA plasmid and assembled by isothermal assembly with
~500bp - 1kb of sequence from near the start and stop codon of the gene to be deleted
and the XhoI/EagI digested pBluescript vector. Primers used to amplify homology arms
are listed in Supplementary File 7(Manage et al 2020). 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). All guide RNA plasmids
were generated by ligating oligos containing the guide RNA sequence into BsaI-digested
pRB1017 (Addgene plasmid # 59936) (Arribere et al. 2014). Guide RNA sequences are
provided in Supplementary File 7(Manage et al 2020). For the introduction of the R159C
mutation in SIMR-1::gfp::3xFLAG, we used an oligo repair template and RNA guide
(Supplementary File 7, Manage et al 2020).
CRISPR injections were performed according to published protocols (Dickinson et al.
2013; 2015; Ward 2015; Arribere et al. 2014; Paix et al. 2015; Dokshin et al. 2018).
CRISPR injection mixes included 10-25 ng/μl repair template, 50 ng/μl guide RNA
plasmid, 50 ng/μl eft-3p::cas9-SV40_NLS::tbb-2 3'UTR (Addgene plasmid # 46168) or
eft-3p::cas9::tbb-2 3'UTR (Addgene plasmid # 61251), 2.5-10 ng/μl GFP or mCherry co-
53
injection markers, and 10 ng/μl hsp-16.1::peel-1 negative selection (pMA122, Addgene
plasmid # 34873). mCherry constructs were injected into USC715: mut-16(cmp3[mut-
16::gfp::3xFLAG + loxP]) I; unc-119(ed3) III. Deletion constructs were injected into
HT1593: unc-119(ed3), except for the hpo-40 deletion construct, which was injected
directly into simr-1(cmp36) I; unc-119(ed3) III. For some strains, floxed Cbr-unc-119(+)
cassettes were excised using eft-3p::Cre (pDD104, Addgene plasmid # 47551)
(Dickinson et al. 2013), however we observed no discernable increase in mCherry-tagged
protein expression after Cbr-unc-119(+) cassette excision. SIMR-1::gfp::3xFLAG and
mKate2::3xMyc::prg-1 was injected into the wild-type strain. For the R159C mutation of
SIMR-1, the injection mix included 0.25 μg/μl Cas9 protein (IDT), 100 ng/μl tracrRNA, 14
ng/μl dpy-10(cn64) crRNA, 42 ng/μl simr-1 crRNA, and 110 ng/μl of each repair template,
and was injected into USC1022(simr-1(cmp112[simr-1::GFP + loxP + 3xFLAG]) I) (Paix
et al. 2015; Dokshin et al. 2018).

Mass Spectrometry
~500,000 synchronized N2 (wild-type) or USC717 (mut-16(cmp3[mut-
16::gfp::3xFLAG + loxP])) adult C. elegans (~68 h at 20°C after L1 arrest) were collected
in IP Buffer (50 mM Tris-Cl pH 7.4, 100 mM KCl, 2.5 mM MgCl2, 0.1% Igapal CA-630, 0.5
mM PMSF (0.5 mM), cOmplete Protease Inhibitor Cocktail (Roche 04693159001), and
RNaseOUT Ribonuclease Inhibitor (ThermoFisher 10777019)), frozen in liquid nitrogen,
and homogenized using a mortar and pestle. After further dilution into IP 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. GFP
54
and FLAG immunoprecipitation was performed at 4°C for 2 hrs using anti-GFP affinity
matrix [RQ2 clone] (MBL International D153-8) and anti-FLAG affinity matrix [M2 clone]
(Sigma-Aldrich A2220), then washed 10 times in immunoprecipitation buffer. 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.

Antibody staining and imaging
Live imaging was conducted by dissecting C. elegans animals in M9 buffer containing
sodium azide and imaging immediately following dissection. For immunofluorescence,
worms were dissected in egg buffer containing 0.1% Tween-20 and fixed in 1%
formaldehyde in egg buffer as described (Phillips et al. 2009). Samples were
immunostained with mouse anti-FLAG (M2, Sigma F1804), rat anti-HA (3F10, Sigma
11867423001), and mouse anti-PGL-1 (DSHB AB 531836). Alexa-Fluor secondary
antibodies were purchased from ThermoFisher Scientific. All worms were dissected as
one-day-old adults (~24 hours after L4). Imaging was performed on a DeltaVision Elite
microscope (GE Healthcare) using 60x N.A. 1.42 oil-immersion objective. When data
stacks were collected, deconvolution was performed using the SoftWoRx package and
presented as maximum intensity projections. Images were pseudocolored using Adobe
Photoshop.
For scoring of apoptotic germ cells, corpses were identified using the bcIs39 (CED-
1::GFP) reporter, which is expressed in gonadal sheath cells and can be observed
surrounding germ cell corpses during engulfment. A minimum of 20 gonads arms were
55
scored per genotype and condition. Information regarding statistical analysis provided in
Supplementary File 8(Manage et al 2020).
Quantification of distance between foci centers was performed in ImageJ according
to published methods (Wan et al. 2018). We imaged pachytene germ cell nuclei from two
animals. Three granules selected from each of four germ cells for a total of 12 granules
per animal. Z stacks were opened using the 3D object counter plugin for ImageJ to collect
the x, y, and z coordinates for the center of each desired foci (Bolte and Cordelières
2006). With these coordinates, distances between the foci centers were calculated using
the distance formula, !(𝑥
!
−𝑥
"
)
!
+ (𝑦
!
−𝑦
"
)
!
+ (𝑧
!
−𝑧
"
)
!
. To account for chromatic
shift between channels, distances were calculated between each pair of channels using
TransFluorospheres streptavidin-labeled microspheres, 0.04 μm (ThermoFisher,
T10711) and these distances were used to correct granule distances.

Protein domain identification
The protein alignment of SIMR-1 with HPO-40 (C. elegans), CJA21107 (C. japonica),
CBN15556 (C. brenneri), and, CRE08315 (C. remanei) was generated using Clustal
Omega and cladogram was made in Evolview V3 (Sievers et al. 2011; Subramanian et
al. 2019). The SIMR-1 protein sequence was input into the HHPred server to identify
remote protein homologs with structural similarity (Söding et al. 2005). A region spanning
amino acids 89-264 of the SIMR-1 protein aligned with extended Tudor domain region of
multiple Tudor domain proteins. A Clustal Omega protein alignment of this putative
extended Tudor domain region of SIMR-1, HPO-40, and their related nematode orthologs
was then generated, and this alignment was entered into the HHpred server to improve
56
sensitivity. The top non-redundant identified proteins, their Protein Data Bank ID code,
and HHpred E-value were H. sapiens TDRD1 (5M9N) – 5.5e-8, M. musculus TDRD1
(4B9X) – 4.3e-7, D. melanogaster Papi/Tdrd2 (5YGB) – 4.5e-7, H. sapiens
SND1/TDRD11 (5M9O) – 7.2e-6, D. melanogaster Tudor (3NTK) – 2.3e-5, H. sapiens
TDRKH/TDRD2 (6B57) – 1.6e-5, D. melanogaster Tudor-SN (2WAC) – 3.2e-5, and B.
mori PAPI (5VQH) – 2.0e-4.

RNAi assays
For RNAi assays, synchronized L1 worms raised at 20°C were fed E. coli expressing
dsRNA against pos-1, lin-29, nhr-23, lir-1, hmr-1, and dpy-13 (Kamath et al. 2003). For
pos-1 and hmr-1, F1 embryos were scored for hatching three to five days after P0 animals
were placed on RNAi bacteria. For lin-29, nhr-23, lir-1, and dpy-13 animals were scored
three days after commencement of feeding RNAi for vulval bursting, larval arrest, larval
arrest, and shorter length (Dumpy), respectively.

Transgenerational fertility and brood size assays
Wild-type and mutant C. elegans strains were maintained at 20°C prior to
temperature-shift experiments. Animals were shifted to 25°C, or back to 20°C, as L1
larvae. For the brood-size assays, 10 L4 animals were picked to individual plates. A single
progeny from each plate was selected and moved to a new plate at L4 stage for the
following generation. If one or more of the animals was sterile, progeny were selected
from one of the replicate plates to maintain the total number of broods scored for each
generation at 10. To score the complete brood, each animal was moved to a fresh plate
57
every day until egg-laying was complete. After allowing the progeny 2-3 days to develop,
the total number of animals on each plate was counted.
For assessment of sperm viability, wild-type and simr-1 mutant males were raised
either at 20°C, a single generation at 25°C, or following 10 generations of growth at 25°C,
and then mated to fog-2 females raised at 20°C. Brood sizes were scored for 10 fog-2
females, each mated to four wild-type or simr-1 mutant males. Males were generated by
heat shock and then maintained as a mating plate at 20°C for multiple generations prior
to beginning temperature-shift experiments. Information regarding statistical analysis
provided in Supplementary File 8(Manage et al 2020).
For assessment of oocyte viability, wild-type and simr-1 mutant hermaphrodites were
raised either at 20°C, a single generation at 25°C, or following 10 generations of growth
at 25°C, and then mated to four pgl-1::gfp males raised at 20°C. Brood sizes were scored
for each of 10 wild-type or simr-1 mutant hermaphrodites, mated to four pgl-1::gfp males.
Only plates where all progeny were GFP positive were scored to ensure that the mating
had occurred. Information regarding statistical analysis provided in Supplementary File
8(Manage et al 2020).

Reestablishing WAGO-class 22G-siRNA production
The mutator pathway was restored to WAGO-class 22G-siRNA-defective animals
according to the crossing scheme in Figure 4B and as previously described (Phillips et al.
2015). The unc-119 mutation was always present in the parental hermaphrodite strain to
allow for unambiguous identification of cross vs. self progeny. F1 animals were singled to
58
individual plates as L4 larvae and scored 2-3 days later for presence or absence of
progeny.

Western blots
For Western blots, proteins were resolved on 4-12% Bis-Tris polyacrylamide gels
(ThermoFisher), transferred to nitrocellulose membranes, and probed with anti-FLAG
1:1,000 [M2 clone] (Sigma-Aldrich F1804), anti-actin 1:10,000 (Abcam ab3280), or anti-
Myc 1:1,000 [9E10 clone] (ThermoFisher 13-2500). Secondary HRP antibodies were
purchased from ThermoFisher.

Small RNA and mRNA library preparation
Small RNAs (18 to 30-nt) were size selected on denaturing 15% polyacrylamide gels
(BioRad 3450091) from total RNA samples. Small RNAs were treated with 5’ RNA
polyphosphatase (Epicentre RP8092H) and ligated to 3’ pre-adenylated adapter with
Truncated T4 RNA ligase (NEB M0373L). Small RNAs were then hybridized to the
reverse transcription primer, ligated to the 5’ adapter with T4 RNA ligase (NEB M0204L),
and reverse transcribed with Superscript III (ThermoFisher 18080-051). Small RNA
libraries were amplified using Q5 High-Fidelity DNA polymerase (NEB M0491L) and size
selected on a 10% polyacrylamide gel (BioRad 3450051).
For mRNA-seq library preparation, nuclease-free H2O was added to 7.5μg of each
RNA sample, extracted from whole animals, to a final volume of 100μL. Samples were
incubated at 65°C for 2 minutes then incubated on ice. The Dynabeads mRNA Purification
Kit (ThermoFisher 61006) was used according to the manufacturer’s protocol. 20μL of
59
Dynabeads was used for each sample. 100ng of each mRNA sample was used to prepare
libraries with the NEBNext Ultra II Directional RNA Library Prep Kit for Illumina (NEB
E7760S) according to the manual, using NEBNext multiplex oligos for Illumina (NEB
E7335S).
Library concentration was determined using the Qubit 1X dsDNA HS Assay kit
(ThermoFisher Q33231) and quality was assessed using the Agilent BioAnalyzer.
Libraries were sequenced on the Illumina NextSeq500 (SE 75-bp reads) platform.

Bioinformatic Analysis
For small RNA libraries, sequences were parsed from adapters using FASTQ/A
Clipper (options: -Q33 -l 17 -c -n -a TGGAATTCTCGGGTGCCAAGG) and quality filtered
using the FASTQ Quality Filter (options: -Q33 -q 27 -p 65) from the FASTX-Toolkit
(http://hannonlab.cshl.edu/fastx_toolkit/), mapped to the C. elegans genome WS258
using Bowtie2 v. 2.2.2 (default parameters) (Langmead and Salzberg 2012), and reads
were assigned to genomic features using FeatureCounts (options: -t exon -g gene_id -O
--fraction –largestOverlap) which is part of the Subread v. 1.5.1 package (Liao et al. 2014;
2013). For all analysis examining total small RNA levels mapping to genes, sequences
were assigned to features in a modified version of the WS258 conical gene set GTF file
where miRNAs and piRNAs were excluded. For mRNA libraries, sequences were parsed
from adapters using Cutadapt v. 1.18 (options: -a
AGATCGGAAGAGCACACGTCTGAACTCCAGTCA -m 17 --nextseq-trim=20 --max-n 2)
(Martin 2011) and mapped to the C. elegans genome WS258 using HISAT2 v. 2.1.0
(options: -k 11) (Kim et al. 2015) and the transcriptome using Salmon v. 0.14.1 (options:
60
-l A --validateMappings) (Patro et al. 2017). Differential expression analysis was done
using DESeq2 v. 1.22.2 (Love et al. 2014). For both small RNA and mRNA-seq libraries,
a two-fold-change cutoff and a DESeq2 adjusted p-value of ≤ 0.05 was required to identify
genes with significant changes in small RNA or mRNA expression. For small RNA-seq
libraries, all genes with differentially-expressed small RNAs also met the requirements of
having at least 10 RPM in either wild-type or mutant libraries. Mutator-target genes,
piRNA-target genes, and ERGO-1-target genes were defined as those whose total
mapped small RNA levels were reduced by at least two-fold in mut-16, prg-1, and ergo-1
mutants compared to wild-type, respectively, with at least 10 RPM in wild-type samples
and a DESeq2 adjusted p-value of ≤ 0.05. PRG-1-independent mutator targets are a
subset of the mutator targets for which the total mapped small RNA levels in prg-1
mutants are either unchanged or increased relative to wild-type. CSR-1 target genes,
ALG-3/4 target genes, spermatogenesis-enriched genes, and oogenesis-enriched genes
were previously described (Lee et al. 2012; Conine et al. 2013; Reinke et al. 2004). All
siRNA target genes are defined as all C. elegans genes with at least 10 RPM in wild-type
or mutant small RNA libraries. Additional data analysis was done using R, Excel, and
custom Python scripts. Venn diagrams were generated using BioVenn (Hulsen et al.
2008) and modified in Adobe Illustrator. Reads per million total reads were plotted along
the WS258 genome using Integrative Genomics Viewer 2.3.90 (Thorvaldsdóttir et al.
2013; Robinson et al. 2011). Sequencing data is summarized in Supplementary File
9(Manage et al 2020).

61
Accession Numbers
High-throughput sequencing data for RNA-sequencing libraries generated during this
study are available through Gene Expression Omnibus (GSE138220 for preliminary simr-
1 small RNA, and simr-1 mRNA sequencing data, GSE134573 for mut-16 small RNA and
mRNA sequencing data, and GSE145217 for prg-1 and ergo-1 small RNA sequencing
data).

Acknowledgements
We thank the members of the Phillips lab for helpful discussions and feedback on the
manuscript. Some strains were provided by the CGC, which is funded by NIH Office of
Research Infrastructure Programs (P40 OD010440), and Shohei Mitani of the National
BioResource Project of Japan. Next generation sequencing was performed by the USC
Molecular Genomics Core, which is supported by award number P30 CA014089 from the
National Cancer Institute, and by the USC Genome Core.

Funding
This work was supported by the National Institute of Health grants K22 CA177897 (to
CMP), R35 GM119656 (to CMP), T32 GM118289 (to DHN), and R35 GM119775 (to
TAM), and the by Deutsche Forschungsgemeinschaft grants KE1888/1-1 and KE1888/1-
2 (to RFK). CMP is a Pew Scholar in the Biomedical Sciences supported by the Pew
Charitable Trusts (www.pewtrusts.org). CJU is funded by the National Science
Foundation Graduate Research Fellowship Program (DGE 1418060) and is a USC
Dornsife-funded Chemistry-Biology Interface trainee.
62

Conflict of interest
The authors declare no competing financial or non-financial interests.
References
Andralojc KM, Campbell AC, Kelly AL, Terrey M, Tanner PC, Gans IM, Senter-Zapata
MJ, Khokhar ES, Updike DL. 2017. ELLI-1, a novel germline protein, modulates RNAi
activity and P-granule accumulation in Caenorhabditis elegans. PLoS Genet 13:
e1006611.
Aravin AA, Sachidanandam R, Girard A, Fejes-Toth K, Hannon GJ. 2007.
Developmentally regulated piRNA clusters implicate MILI in transposon control.
Science 316: 744–747.
Arribere JA, Bell RT, Fu BXH, Artiles KL, Hartman PS, Fire AZ. 2014. Efficient Marker-
Free Recovery of Custom Genetic Modifications with CRISPR/Cas9 in
Caenorhabditis elegans. Genetics 198: 837–846.
Ashe A, Sapetschnig A, Weick E-M, Mitchell J, Bagijn MP, Cording AC, Doebley A-L,
Goldstein LD, Lehrbach NJ, Le Pen J, et al. 2012. piRNAs Can Trigger a
Multigenerational Epigenetic Memory in the Germline of C. elegans. Cell 150: 88–
99.
Bagijn MP, Goldstein LD, Sapetschnig A, Weick E-M, Bouasker S, Lehrbach NJ, Simard
MJ, Miska EA. 2012. Function, Targets, and Evolution of Caenorhabditis elegans
piRNAs. Science 337: 574–578.
Barucci G, Cornes E, Singh M, Li B, Ugolini M, Samolygo A, Didier C, Dingli F, Loew D,
Quarato P, et al. 2020. Small-RNA-mediated transgenerational silencing of histone
genes impairs fertility in piRNA mutants. Nat Cell Biol 22: 235–245.
Batista PJ, Ruby JG, Claycomb JM, Chiang R, Fahlgren N, Kasschau KD, Chaves DA,
Gu W, Vasale JJ, Duan S, et al. 2008. PRG-1 and 21U-RNAs interact to form the
piRNA complex required for fertility in C. elegans. Mol Cell 31: 67–78.
Bessereau J-L. 2006. Transposons in C. elegans. WormBook 1–13.
Bélicard T, Jareosettasin P, Sarkies P. 2018. The piRNA pathway responds to
environmental signals to establish intergenerational adaptation to stress. BMC Biol
16: 103.
Billi AC, Alessi AF, Khivansara V, Han T, Freeberg M, Mitani S, Kim JK. 2012. The
Caenorhabditis elegans HEN1 Ortholog, HENN-1, Methylates and Stabilizes Select
Subclasses of Germline Small RNAs. PLoS Genet 8: e1002617.
63
Bolte S, Cordelières FP. 2006. A guided tour into subcellular colocalization analysis in
light microscopy. J Microsc 224: 213–232.
Boswell RE, Mahowald AP. 1985. tudor, a gene required for assembly of the germ
plasm in Drosophila melanogaster. Cell 43: 97–104.
Brangwynne CP, Eckmann CR, Courson DS, Rybarska A, Hoege C, Gharakhani J,
Jülicher F, Hyman AA. 2009. Germline P granules are liquid droplets that localize by
controlled dissolution/condensation. Science 324: 1729–1732.
Brennecke J, Aravin AA, Stark A, Dus M, Kellis M, Sachidanandam R, Hannon GJ.
2007. Discrete small RNA-generating loci as master regulators of transposon
activity in Drosophila. Cell 128: 1089–1103.
Brenner S. 1974. The genetics of Caenorhabditis elegans. Genetics 77: 71–94.
Brownlie JC, Whyard S. 2004. CemaT1 is an active transposon within the
Caenorhabditis elegans genome. Gene 338: 55–64.
Buckley BA, Burkhart KB, Gu SG, Spracklin G, Kershner A, Fritz H, Kimble J, Fire A,
Kennedy S. 2012. A nuclear Argonaute promotes multigenerational epigenetic
inheritance and germline immortality. Nature 489: 447–451.
Callebaut I, Mornon JP. 1997. The human EBNA-2 coactivator p100: multidomain
organization and relationship to the staphylococcal nuclease fold and to the tudor
protein involved in Drosophila melanogaster development. Biochem J 321 ( Pt 1):
125–132.
Chen C, Nott TJ, Jin J, Pawson T. 2011. Deciphering arginine methylation: Tudor tells
the tale. Nat Rev Mol Cell Biol 12: 629–642.
Claycomb JM. 2014. Ancient endo-siRNA pathways reveal new tricks. Curr Biol 24:
R703–15.
Claycomb JM, Batista PJ, Pang KM, Gu W, Vasale JJ, van Wolfswinkel JC, Chaves DA,
Shirayama M, Mitani S, Ketting RF, et al. 2009. The Argonaute CSR-1 and its 22G-
RNA cofactors are required for holocentric chromosome segregation. Cell 139: 123–
134.
Collins J, Forbes E, Anderson P. 1989. The Tc3 family of transposable genetic
elements in Caenorhabditis elegans. Genetics 121: 47–55.
Collins JJ, Anderson P. 1994. The Tc5 family of transposable elements in
Caenorhabditis elegans. Genetics 137: 771–781.
Conine CC, Moresco JJ, Gu W, Shirayama M, Conte D, Yates JR, Mello CC. 2013.
Argonautes Promote Male Fertility and Provide a Paternal Memory of Germline
Gene Expression in C. elegans. Cell 155: 1532–1544.
64
Cordeiro Rodrigues RJ, de Jesus Domingues AM, Hellmann S, Dietz S, de Albuquerque
BFM, Renz C, Ulrich HD, Sarkies P, Butter F, Ketting RF. 2019. PETISCO is a novel
protein complex required for 21U RNA biogenesis and embryonic viability. Genes
Dev.
Corrêa RL, Steiner FA, Berezikov E, Ketting RF. 2010. MicroRNA-directed siRNA
biogenesis in Caenorhabditis elegans. PLoS Genet 6: e1000903.
Das PP, Bagijn MP, Goldstein LD, Woolford JR, Lehrbach NJ, Sapetschnig A, Buhecha
HR, Gilchrist MJ, Howe KL, Stark R, et al. 2008. Piwi and piRNAs act upstream of
an endogenous siRNA pathway to suppress Tc3 transposon mobility in the
Caenorhabditis elegans germline. Mol Cell 31: 79–90.
de Albuquerque BFM, Luteijn MJ, Cordeiro Rodrigues RJ, van Bergeijk P, Waaijers S,
Kaaij LJT, Klein H, Boxem M, Ketting RF. 2014. PID-1 is a novel factor that operates
during 21U-RNA biogenesis in Caenorhabditis elegans. Genes Dev 28: 683–688.
de Albuquerque BFM, Placentino M, Ketting RF. 2015. Maternal piRNAs Are Essential
for Germline Development following De Novo Establishment of Endo-siRNAs in
Caenorhabditis elegans. Dev Cell 34: 448–456.
Dickinson DJ, Pani AM, Heppert JK, Higgins CD, Goldstein B. 2015. Streamlined
Genome Engineering with a Self-Excising Drug Selection Cassette. Genetics 200:
1035–1049.
Dickinson DJ, Ward JD, Reiner DJ, Goldstein B. 2013. Engineering the Caenorhabditis
elegans genome using Cas9-triggered homologous recombination. Nat Methods 10:
1028–1034.
Dokshin GA, Ghanta KS, Piscopo KM, Mello CC. 2018. Robust Genome Editing with
Short Single-Stranded and Long, Partially Single-Stranded DNA Donors in
Caenorhabditis elegans. Genetics 210: 781–787.
Eide D, Anderson P. 1985. Transposition of Tc1 in the nematode Caenorhabditis
elegans. Proc Natl Acad Sci USA 82: 1756–1760.
Fischer SEJ, Butler MD, Pan Q, Ruvkun G. 2008. Trans-splicing in C. elegans
generates the negative RNAi regulator ERI-6/7. Nature 455: 491–496.
Fischer SEJ, Montgomery TA, Zhang C, Fahlgren N, Breen PC, Hwang A, Sullivan CM,
Carrington JC, Ruvkun G. 2011. The ERI-6/7 Helicase Acts at the First Stage of an
siRNA Amplification Pathway That Targets Recent Gene Duplications. PLoS Genet
7: e1002369.
Friberg A, Corsini L, Mourão A, Sattler M. 2009. Structure and ligand binding of the
extended Tudor domain of D. melanogaster Tudor-SN. J Mol Biol 387: 921–934.

65
Friesen WJ, Massenet S, Paushkin S, Wyce A, Dreyfuss G. 2001. SMN, the product of
the spinal muscular atrophy gene, binds preferentially to dimethylarginine-containing
protein targets. Mol Cell 7: 1111–1117.
Gartner A, Boag PR, Blackwell TK. 2008. Germline survival and apoptosis. WormBook
1–20.
Gartner A, Milstein S, Ahmed S, Hodgkin J, Hengartner MO. 2000. A conserved
checkpoint pathway mediates DNA damage-induced apoptosis and cell cycle arrest
in C. elegans. Mol Cell 5: 435–443.
Gibson DG, Young L, Chuang R-Y, Venter JC, Hutchison CA, Smith HO. 2009.
Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat
Methods 6: 343–345.
Gu W, Shirayama M, Conte D, Vasale J, Batista PJ, Claycomb JM, Moresco JJ,
Youngman EM, Keys J, Stoltz MJ, et al. 2009. Distinct argonaute-mediated 22G-
RNA pathways direct genome surveillance in the C. elegans germline. Mol Cell 36:
231–244.
Gumienny TL, Lambie E, Hartwieg E, Horvitz HR, Hengartner MO. 1999. Genetic
control of programmed cell death in the Caenorhabditis elegans hermaphrodite
germline. Development 126: 1011–1022.
Gunawardane LS, Saito K, Nishida KM, Miyoshi K, Kawamura Y, Nagami T, Siomi H,
Siomi MC. 2007. A slicer-mediated mechanism for repeat-associated siRNA 5' end
formation in Drosophila. Science 315: 1587–1590.
Han W, Sundaram P, Kenjale H, Grantham J, Timmons L. 2008. The Caenorhabditis
elegans rsd-2 and rsd-6 genes are required for chromosome functions during
exposure to unfavorable environments. Genetics 178: 1875–1893.
Handler D, Olivieri D, Novatchkova M, Gruber FS, Meixner K, Mechtler K, Stark A,
Sachidanandam R, Brennecke J. 2011. A systematic analysis of Drosophila TUDOR
domain-containing proteins identifies Vreteno and the Tdrd12 family as essential
primary piRNA pathway factors. EMBO J 30: 3977–3993.
Houwing S, Kamminga LM, Berezikov E, Cronembold D, Girard A, van den Elst H,
Filippov DV, Blaser H, Raz E, Moens CB, et al. 2007. A role for Piwi and piRNAs in
germ cell maintenance and transposon silencing in Zebrafish. Cell 129: 69–82.
Hulsen T, de Vlieg J, Alkema W. 2008. BioVenn - a web application for the comparison
and visualization of biological lists using area-proportional Venn diagrams. BMC
Genomics 9: 488.
Hutvagner G, Simard MJ. 2008. Argonaute proteins: key players in RNA silencing. Nat
Rev Mol Cell Biol 9: 22–32.
66
Kamath RS, Fraser AG, Dong Y, Poulin G, Durbin R, Gotta M, Kanapin A, Le Bot N,
Moreno S, Sohrmann M, et al. 2003. Systematic functional analysis of the
Caenorhabditis elegans genome using RNAi. Nature 421: 231–237.
Kamminga LM, van Wolfswinkel JC, Luteijn MJ, Kaaij LJT, Bagijn MP, Sapetschnig A,
Miska EA, Berezikov E, Ketting RF. 2012. Differential Impact of the HEN1 Homolog
HENN-1 on 21U and 26G RNAs in the Germline of Caenorhabditis elegans. PLoS
Genet 8: e1002702.
Kasper DM, Wang G, Gardner KE, Johnstone TG, Reinke V. 2014. The C. elegans
SNAPc Component SNPC-4 Coats piRNA Domains and Is Globally Required for
piRNA Abundance. Dev Cell 31: 145–158.
Ketting RF. 2011. The many faces of RNAi. Dev Cell 20: 148–161.
Ketting RF, Haverkamp TH, van Luenen HG, Plasterk RHA. 1999. Mut-7 of C. elegans,
required for transposon silencing and RNA interference, is a homolog of Werner
syndrome helicase and RNaseD. Cell 99: 133–141.
Ketting RF, Plasterk RHA. 2000. A genetic link between co-suppression and RNA
interference in C. elegans. Nature 404: 296–298.
Kim D, Langmead B, Salzberg SL. 2015. HISAT: a fast spliced aligner with low memory
requirements. Nat Methods 12: 357–360.
Kirino Y, Kim N, de Planell-Saguer M, Khandros E, Chiorean S, Klein PS, Rigoutsos I,
Jongens TA, Mourelatos Z. 2009. Arginine methylation of Piwi proteins catalysed by
dPRMT5 is required for Ago3 and Aub stability. Nat Cell Biol 11: 652–658.
Langmead B, Salzberg SL. 2012. Fast gapped-read alignment with Bowtie 2. Nat
Methods 9: 357–359.
Lee H-C, Gu W, Shirayama M, Youngman E, Conte D, Mello CC. 2012. C. elegans
piRNAs Mediate the Genome-wide Surveillance of Germline Transcripts. Cell 150:
78–87.
Levitt A, Emmons SW. 1989. The Tc2 transposon in Caenorhabditis elegans. Proc Natl
Acad Sci USA 86: 3232–3236.
Li W, Shaw JE. 1993. A variant Tc4 transposable element in the nematode C. elegans
could encode a novel protein. Nucleic Acids Res 21: 59–67.
Liao Y, Smyth GK, Shi W. 2014. featureCounts: an efficient general purpose program
for assigning sequence reads to genomic features. Bioinformatics 30: 923–930.
Liao Y, Smyth GK, Shi W. 2013. The Subread aligner: fast, accurate and scalable read
mapping by seed-and-vote. Nucleic Acids Res 41: e108.
67
Liu H, Wang J-YS, Huang Y, Li Z, Gong W, Lehmann R, Xu R-M. 2010a. Structural
basis for methylarginine-dependent recognition of Aubergine by Tudor. Genes Dev
24: 1876–1881.
Liu K, Chen C, Guo Y, Lam R, Bian C, Xu C, Zhao DY, Jin J, MacKenzie F, Pawson T,
et al. 2010b. Structural basis for recognition of arginine methylated Piwi proteins by
the extended Tudor domain. Proc Natl Acad Sci USA 107: 18398–18403.
Love MI, Huber W, Anders S. 2014. Moderated estimation of fold change and
dispersion for RNA-seq data with DESeq2. Genome Biol 15: 550.
Luteijn MJ, van Bergeijk P, Kaaij LJT, Almeida MV, Roovers EF, Berezikov E, Ketting
RF. 2012. Extremely stable Piwi-induced gene silencing in Caenorhabditis elegans.
EMBO J 31: 3422–3430.
Marchler-Bauer A, Lu S, Anderson JB, Chitsaz F, Derbyshire MK, DeWeese-Scott C,
Fong JH, Geer LY, Geer RC, Gonzales NR, et al. 2011. CDD: a Conserved Domain
Database for the functional annotation of proteins. Nucleic Acids Res 39: D225–9.
Martin M. 2011. Cutadapt removes adapter sequences from high-throughput
sequencing reads. EMBnet 17: 10–12.
Mathioudakis N, Palencia A, Kadlec J, Round A, Tripsianes K, Sattler M, Pillai RS,
Cusack S. 2012. The multiple Tudor domain-containing protein TDRD1 is a
molecular scaffold for mouse Piwi proteins and piRNA biogenesis factors. RNA 18:
2056–2072.
McMurchy AN, Stempor P, Gaarenstroom T, Wysolmerski B, Dong Y, Aussianikava D,
Appert A, Huang N, Kolasinska-Zwierz P, Sapetschnig A, et al. 2017. A team of
heterochromatin factors collaborates with small RNA pathways to combat repetitive
elements and germline stress. Elife 6.
Montgomery TA, Rim Y-S, Zhang C, Dowen RH, Phillips CM, Fischer SEJ, Ruvkun G.
2012. PIWI Associated siRNAs and piRNAs Specifically Require the Caenorhabditis
elegans HEN1 Ortholog henn-1. PLoS Genet 8: e1002616.
Ni JZ, Kalinava N, Chen E, Huang A, Trinh T, Gu SG. 2016. A transgenerational role of
the germline nuclear RNAi pathway in repressing heat stress-induced transcriptional
activation in C. elegans. Epigenetics Chromatin 9: 3.
Nishida KM, Okada TN, Kawamura T, Mituyama T, Kawamura Y, Inagaki S, Huang H,
Chen D, Kodama T, Siomi H, et al. 2009. Functional involvement of Tudor and
dPRMT5 in the piRNA processing pathway in Drosophila germlines. EMBO J 28:
3820–3831.
Paix A, Folkmann A, Rasoloson D, Seydoux G. 2015. High Efficiency, Homology-
Directed Genome Editing in Caenorhabditis elegans Using CRISPR-Cas9
Ribonucleoprotein Complexes. Genetics 201: 47–54.
68
Pak J, Fire A. 2007. Distinct populations of primary and secondary effectors during
RNAi in C. elegans. Science 315: 241–244.
Patro R, Duggal G, Love MI, Irizarry RA, Kingsford C. 2017. Salmon provides fast and
bias-aware quantification of transcript expression. Nat Methods 14: 417–419.
Pek JW, Anand A, Kai T. 2012. Tudor domain proteins in development. Development
139: 2255–2266.
Pettitt J, Crombie C, Schumperli D, Muller B. 2002. The Caenorhabditis elegans histone
hairpin-binding protein is required for core histone gene expression and is essential
for embryonic and postembryonic cell division. J Cell Sci 115: 857–866.
Phillips CM, Brown KC, Montgomery BE, Ruvkun G, Montgomery TA. 2015. piRNAs
and piRNA-Dependent siRNAs Protect Conserved and Essential C. elegans Genes
from Misrouting into the RNAi Pathway. Dev Cell 34: 457–465.
Phillips CM, McDonald KL, Dernburg AF. 2009. Cytological analysis of meiosis in
Caenorhabditis elegans. Methods Mol Biol 558: 171–195.
Phillips CM, Montgomery BE, Breen PC, Roovers EF, Rim Y-S, Ohsumi TK, Newman
MA, van Wolfswinkel JC, Ketting RF, Ruvkun G, et al. 2014. MUT-14 and SMUT-1
DEAD Box RNA Helicases Have Overlapping Roles in Germline RNAi and
Endogenous siRNA Formation. Curr Biol 24: 839–844.
Phillips CM, Montgomery TA, Breen PC, Ruvkun G. 2012. MUT-16 promotes formation
of perinuclear Mutator foci required for RNA silencing in the C. elegans germline.
Genes Dev 26: 1433–1444.
Ponting CP. 1997. Tudor domains in proteins that interact with RNA. Trends Biochem
Sci 22: 51–52.
Reed KJ, Svendsen JM, Brown KC, Montgomery BE, Marks TN, Vijayasarathy T,
Parker DM, Nishimura EO, Updike DL, Montgomery TA. 2020. Widespread roles for
piRNAs and WAGO-class siRNAs in shaping the germline transcriptome of
Caenorhabditis elegans. Nucleic Acids Res 48: 1811–1827.
Reinke V, Gil IS, Ward S, Kazmer K. 2004. Genome-wide germline-enriched and sex-
biased expression profiles in Caenorhabditis elegans. Development 131: 311–323.
Ren R, Liu H, Wang W, Wang M, Yang N, Dong Y-H, Gong W, Lehmann R, Xu R-M.
2014. Structure and domain organization of Drosophila Tudor. Cell Res 24: 1146–
1149.
Reuter M, Chuma S, Tanaka T, Franz T, Stark A, Pillai RS. 2009. Loss of the Mili-
interacting Tudor domain-containing protein-1 activates transposons and alters the
Mili-associated small RNA profile. Nat Struct Mol Biol 16: 639–646.
69
Rezsohazy R, van Luenen HG, Durbin RM, Plasterk RHA. 1997. Tc7, a Tc1-hitch hiking
transposon in Caenorhabditis elegans. Nucleic Acids Res 25: 4048–4054.
Robert VJ, Sijen T, van Wolfswinkel J, Plasterk RHA. 2005. Chromatin and RNAi factors
protect the C. elegans germline against repetitive sequences. Genes Dev 19: 782–
787.
Robinson JT, Thorvaldsdóttir H, Winckler W, Guttman M, Lander ES, Getz G, Mesirov
JP. 2011. Integrative genomics viewer. Nat Biotechnol 29: 24–26.
Rogers AK, Phillips CM. 2020. RNAi pathways repress reprogramming of C. elegans
germ cells during heat stress. Nucleic Acids Res.
Sakaguchi A, Sarkies P, Simon M, Doebley A-L, Goldstein LD, Hedges A, Ikegami K,
Alvares SM, Yang L, LaRocque JR, et al. 2014. Caenorhabditis elegans RSD-2 and
RSD-6 promote germ cell immortality by maintaining small interfering RNA
populations. Proc Natl Acad Sci USA 111: E4323–31.
Schumacher B, Hanazawa M, Lee M-H, Nayak S, Volkmann K, Hofmann ER, Hofmann
R, Hengartner M, Schedl T, Gartner A. 2005. Translational repression of C. elegans
p53 by GLD-1 regulates DNA damage-induced apoptosis. Cell 120: 357–368.
Shen E-Z, Chen H, Ozturk AR, Tu S, Shirayama M, Tang W, Ding Y-H, Dai S-Y, Weng
Z, Mello CC. 2018. Identification of piRNA Binding Sites Reveals the Argonaute
Regulatory Landscape of the C. elegans Germline. Cell 172: 937–951.e18.
Shirayama M, Seth M, Lee H-C, Gu W, Ishidate T, Conte D, Mello CC. 2012. piRNAs
Initiate an Epigenetic Memory of Nonself RNA in the C. elegans Germline. Cell 150:
65–77.
Shirayama M, Stanney W, Gu W, Seth M, Mello CC. 2014. The Vasa Homolog RDE-12
Engages Target mRNA and Multiple Argonaute Proteins to Promote RNAi in
C. elegans. Curr Biol 24: 845–851.
Sievers F, Wilm A, Dineen D, Gibson TJ, Karplus K, Li W, Lopez R, McWilliam H,
Remmert M, Söding J, et al. 2011. Fast, scalable generation of high-quality protein
multiple sequence alignments using Clustal Omega. Mol Syst Biol 7: 539.
Sijen T, Steiner FA, Thijssen KL, Plasterk RHA. 2007. Secondary siRNAs result from
unprimed RNA synthesis and form a distinct class. Science 315: 244–247.
Simon M, Sarkies P, Ikegami K, Doebley A-L, Goldstein LD, Mitchell J, Sakaguchi A,
Miska EA, Ahmed S. 2014. Reduced Insulin/IGF-1 Signaling Restores Germ Cell
Immortality to Caenorhabditis elegans Piwi Mutants. Cell Rep 7: 762–773.
Söding J, Biegert A, Lupas AN. 2005. The HHpred interactive server for protein
homology detection and structure prediction. Nucleic Acids Res 33: W244–8.
70
Spracklin G, Fields B, Wan G, Becker D, Wallig A, Shukla A, Kennedy S. 2017. The
RNAi Inheritance Machinery of Caenorhabditis elegans. Genetics 206: 1403–1416.
Subramanian B, Gao S, Lercher MJ, Hu S, Chen W-H. 2019. Evolview v3: a webserver
for visualization, annotation, and management of phylogenetic trees. Nucleic Acids
Res 47: W270–W275.
Tabara H, Sarkissian M, Kelly WG, Fleenor J, Grishok A, Timmons L, Fire A, Mello CC.
1999. The rde-1 gene, RNA interference, and transposon silencing in C. elegans.
Cell 99: 123–132.
Tang W, Tu S, Lee H-C, Weng Z, Mello CC. 2016. The RNase PARN-1 Trims piRNA 3'
Ends to Promote Transcriptome Surveillance in C. elegans. Cell 164: 974–984.
Thorvaldsdóttir H, Robinson JT, Mesirov JP. 2013. Integrative Genomics Viewer (IGV):
high-performance genomics data visualization and exploration. Brief Bioinformatics
14: 178–192.
Tijsterman M, May RC, Simmer F, Okihara KL, Plasterk RHA. 2004. Genes required for
systemic RNA interference in Caenorhabditis elegans. Curr Biol 14: 111–116.
Tsai H-Y, Chen C-CG, Conte D, Moresco JJ, Chaves DA, Mitani S, Yates JR, Tsai M-D,
Mello CC. 2015. A Ribonuclease Coordinates siRNA Amplification and mRNA
Cleavage during RNAi. Cell 160: 407–419.
Uebel CJ, Anderson DC, Mandarino LM, Manage KI, Aynaszyan S, Phillips CM. 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.
Vagin VV, Wohlschlegel JA, Qu J, Jonsson Z, Huang X, Chuma S, Girard A,
Sachidanandam R, Hannon GJ, Aravin AA. 2009. Proteomic analysis of murine Piwi
proteins reveals a role for arginine methylation in specifying interaction with Tudor
family members. Genes Dev 23: 1749–1762.
Vastenhouw NL, Fischer SEJ, Robert VJ, Thijssen KL, Fraser AG, Kamath RS, Ahringer
J, Plasterk RHA. 2003. A genome-wide screen identifies 27 genes involved in
transposon silencing in C. elegans. Curr Biol 13: 1311–1316.
Wallis DC, Nguyen DAH, Uebel CJ, Phillips CM. 2019. Visualization and Quantification
of Transposon Activity in Caenorhabditis elegans RNAi Pathway Mutants. G3
(Bethesda) 9: 3825–3832.
Wan G, Fields BD, Spracklin G, Shukla A, Phillips CM, Kennedy S. 2018.
Spatiotemporal regulation of liquid-like condensates in epigenetic inheritance.
Nature 557: 679–683.

71
Wang G, Reinke V. 2008. A C. elegans Piwi, PRG-1, regulates 21U-RNAs during
spermatogenesis. Curr Biol 18: 861–867.
Ward JD. 2015. Rapid and precise engineering of the Caenorhabditis elegans genome
with lethal mutation co-conversion and inactivation of NHEJ repair. Genetics 199:
363–377.
Webster A, Li S, Hur JK, Wachsmuth M, Bois JS, Perkins EM, Patel DJ, Aravin AA.
2015. Aub and Ago3 Are Recruited to Nuage through Two Mechanisms to Form a
Ping-Pong Complex Assembled by Krimper. Mol Cell 59: 564–575.
Wedeles CJ, Wu MZ, Claycomb JM. 2013. Protection of germline gene expression by
the C. elegans Argonaute CSR-1. Dev Cell 27: 664–671.
Weick E-M, Miska EA. 2014. piRNAs: from biogenesis to function. Development 141:
3458–3471.
Weick E-M, Sarkies P, Silva N, Chen RA, Moss SMM, Cording AC, Ahringer J,
Martinez-Perez E, Miska EA. 2014. PRDE-1 is a nuclear factor essential for the
biogenesis of Ruby motif-dependent piRNAs in C. elegans. Genes Dev 28: 783–
796.
Wu W-S, Brown JS, Chen T-T, Chu Y-H, Huang W-C, Tu S, Lee H-C. 2019.
piRTarBase: a database of piRNA targeting sites and their roles in gene regulation.
Nucleic Acids Res 47: D181–D187.
Wu W-S, Huang W-C, Brown JS, Zhang D, Song X, Chen H, Tu S, Weng Z, Lee H-C.
2018. pirScan: a webserver to predict piRNA targeting sites and to avoid transgene
silencing in C. elegans. Nucleic Acids Res 46: W43–W48.
Yang H, Vallandingham J, Shiu P, Li H, Hunter CP, Mak HY. 2014. The DEAD Box
Helicase RDE-12 Promotes Amplification of RNAi in Cytoplasmic Foci in C. elegans.
Curr Biol 24: 832–838.
Yang H, Zhang Y, Vallandingham J, Li H, Florens L, Mak HY. 2012. The RDE-10/RDE-
11 complex triggers RNAi-induced mRNA degradation by association with target
mRNA in C. elegans. Genes Dev 26: 846–856.
Yigit E, Batista PJ, Bei Y, Pang KM, Chen C-CG, Tolia NH, Joshua-Tor L, Mitani S,
Simard MJ, Mello CC. 2006. Analysis of the C. elegans Argonaute family reveals
that distinct Argonautes act sequentially during RNAi. Cell 127: 747–757.
Yuan JY, Finney M, Tsung N, Horvitz HR. 1991. Tc4, a Caenorhabditis elegans
transposable element with an unusual fold-back structure. Proc Natl Acad Sci USA
88: 3334–3338.

72
Zeng C, Weng C, Wang X, Yan Y-H, Li W-J, Xu D, Hong M, Liao S, Dong M-Q, Feng X,
et al. 2019. Functional Proteomics Identifies a PICS Complex Required for piRNA
Maturation and Chromosome Segregation. Cell Rep 27: 3561–3572.e3.
Zhang C, Montgomery TA, Fischer SEJ, Garcia SMDA, Riedel CG, Fahlgren N, Sullivan
CM, Carrington JC, Ruvkun G. 2012. The Caenorhabditis elegans RDE-10/RDE-11
Complex Regulates RNAi by Promoting Secondary siRNA Amplification. Curr Biol
22: 881–890.
Zhang C, Montgomery TA, Gabel HW, Fischer SEJ, Phillips CM, Fahlgren N, Sullivan
CM, Carrington JC, Ruvkun G. 2011. mut-16 and other mutator class genes
modulate 22G and 26G siRNA pathways in Caenorhabditis elegans. Proc Natl Acad
Sci USA 108: 1201–1208.
Zhang D, Tu S, Stubna M, Wu W-S, Huang W-C, Weng Z, Lee H-C. 2018a. The piRNA
targeting rules and the resistance to piRNA silencing in endogenous genes. Science
359: 587–592.
Zhang H, Liu K, Izumi N, Huang H, Ding D, Ni Z, Sidhu SS, Chen C, Tomari Y, Min J.
2017. Structural basis for arginine methylation-independent recognition of PIWIL1
by TDRD2. Proc Natl Acad Sci USA 114: 12483–12488.
Zhang Y, Liu W, Li R, Gu J, Wu P, Peng C, Ma J, Wu L, Yu Y, Huang Y. 2018b.
Structural insights into the sequence-specific recognition of Piwi by Drosophila Papi.
Proc Natl Acad Sci USA 115: 3374–3379.

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Figures and Figure Legends

Figure 1. SIMR-1 is a perinuclear-localized Tudor domain protein.
(A) Proteins identified by IP-mass spec of MUT-16::GFP::3xFLAG but not wild-type animals. The percent coverage and
total number of peptides captured are indicated for each MUT-16-associated protein. See Supplementary File
2(Manage et al 2020) for complete list of immunoprecipitated proteins. (B) Live imaging of SIMR-1::mCherry
demonstrate that it is adjacent to or colocalizes with MUT-16::GFP foci. Scale bars, 5μm. (C) Cladogram representing
the relationship between SIMR-1 and related proteins CJA21107 (C. japonica), CBN15556 (C. brenneri), CRE08315
(C. remanei), and HPO-40 (C. elegans). The protein alignment was generated using Clustal Omega and cladogram
was made in Evolview V3. (D) Live imaging of SIMR-1::mCherry in a mut-16 mutant and MUT-16::GFP in a simr-1;
hpo-40 double mutant indicate that mut-16 is not required for SIMR-1 foci formation, nor are simr-1 and hpo-40 required
for Mutator foci formation. Scale bars, 5μm. (E) Alignment of Tudor domain region generated by Clustal Omega of
SIMR-1, HPO-40, their related nematode orthologs, and the eight most significant hits from HHpred server (see
Methods). The four aromatic residues that constitute the aromatic cage are highlighted in blue and the absolutely
conserved arginine and aspartate residues characteristic of extended Tudor domains are highlighted in red. The
location of the simr-1[R159C] mutation is marked with an asterisk. Cel - C. elegans, Cre – C. remanei, Cbn – C.
brenneri, Cja – C. japonica, Dme – D. melanogaster, Hsa – H. sapiens, Mmu – M. musculus, and Bmo – B. mori.

74

Figure 1–figure supplement 1. Identification and localization of MUT-16-associated proteins.
(A) Proteins identified by IP-mass spec of MUT-16::GFP::3xFLAG that are enriched at least four-fold in MUT-16
immunoprecipitations relative to control immunoprecipitations. See Supplementary File 2(Manage et al 2020) for
complete list of immunoprecipitated proteins. The percent coverage and total number of peptides captured are indicated
for each MUT-16-associated protein. (B) Live imaging of C33G3.6::mCherry, F37C4.5::mCherry, HGO-1::mCherry,
MATH-33::mCherry, and Y57G11C.3::mCherry. MUT-16::GFP is shown for reference. (C-D) Live imaging of RSD-
2::mCherry (C) and HPO-40::mCherry (D) demonstrates that both proteins are adjacent to or colocalize with MUT-
16::GFP foci. Scale bars, 5μm.

HPO-40::mCherry
MUT-16::GFP
merge
A
MUT-16 control MUT-16 control
RSD-2/F52G2.2 Unknown function 25.63% (32) 2.2% (2) 35.18% (42) 0% (0)
WAGO-1/R06C7.1 WAGO Argonaute 17.14% (14) 3.92% (3) 13.86% (14) 0% (0)
MATH-33/H19N07.2 Ubiquitin C-terminal hydrolase 11.28% (11) 1.15% (1) 13.13% (11) 0% (0)
Protein Description FLAG IP GFP IP
% Coverage (Peptide Counts)
MUT-16::GFP
C33G3.6::mCherry
Y57G11C.3::mCherry
HGO-1::mCherry F37C4.5::mCherry
MATH-33::mCherry
B
RSD-2::mCherry
merge
MUT-16::GFP
C D
75

Figure 2. Small RNA-related phenotypes associated with deletions in MUT-16-associated proteins.
(A) Animals carrying deletions for each previously-uncharacterized gene identified in the MUT-16 IP-mass spec
experiment were assayed for their ability to respond to somatic (nhr-23 or lin-29) or germline (pos-1) RNAi. “+” indicates
wild-type response and “-” indicates RNAi-defective response. (B) Worms carrying deletions for hpo-40 single mutants
or simr-1; hpo-40 double mutants were assayed for their ability to respond to somatic (nhr-23 or lin-29) or germline
(pos-1) RNAi as described in (A). (C) Box plot displaying total small RNA levels targeting mutator-target genes in the
indicated mutant strains relative to wild-type animals. (D,E) Scatter plots display small RNA reads per million total reads
mapping to mutator-target genes (D) and CSR-1-class genes (E) in wild-type and simr-1 mutants. Genes for which
log2(fold change small RNA abundance) ≥ 1 are colored dark red and genes for which log2(fold change small RNA
abundance) ≤ -1 are colored light blue.
simr-1
hgo-1
math-33
mut-16
nyn-1; nyn-2
rsd-2
wago-1
Known mutator
complex genes
RNAi-
related
Unknown
small RNAs targeting mutator-class genes
rde-8
log
(fold change small RNA abundance)
(mutant/wild-type)
A C
wild-type (RPM)
simr-1 (RPM)
small RNAs targeting mutator-class genes
nhr-23 lin-29 pos-1
wild-type
mut-16
nyn-1; nyn-2
rde-8
rsd-2
wago-1
hgo-1
math-33
simr-1
+
gene
+ +
+ + +
+ + +
+ + +
+ + +
+ + +
+ + +
+ + +
- - -
- - -
- - -
- - -
nhr-23 lin-29 pos-1
hpo-40
simr-1; hpo-40
gene
+ + +
+ + +
B
D E
simr-1 (RPM)
small RNAs targeting wild-type (RPM)
log
log
log
log

76

Figure 2–figure supplement 1. Mutator-class small RNAs are reduced in simr-1 but not hpo-40
mutants.
(A) Box plot displaying total small RNAs levels targeting mutator-target genes in the indicated mutant strains relative to
wild-type animals. (B) Scatter plots display small RNA reads per million total reads mapping to mutator-target genes in
wild-type and hpo-40 single mutants (left) or simr-1; hpo-40 double mutants (right). Genes for which log2(fold change
small RNA abundance) ≥ 1 are colored dark red and genes for which log2(fold change small RNA abundance) ≤ -1 are
colored light blue.
simr-1
simr-1; hpo-40
hpo-40
small RNAs targeting
mutator-class genes
A B
wild-type (RPM)
hpo-40 (RPM)
small RNAs targeting mutator-class genes
wild-type (RPM)
simr-1; hpo-40 (RPM)
small RNAs targeting mutator-class genes
log
(fold change small RNA abundance)
(mutant/wild-type)
log
log
log
log

77

Figure 3. simr-1 mutants have a transgenerational fertility defect at elevated temperature.
(A) Brood size was scored for a single generation at 20°C, followed by 11 generations at 25°C, demonstrating that
simr-1 mutants become progressively sterile at 25°C. 10 broods were scored for each genotype at each generation.
(B) Brood sizes for simr-1 mutant and wild-type animals were scored for 10 generations after returning to 20°C, following
10 generations at 25°C, demonstrating restoration of fertility at permissive temperature. 10 broods were scored for
each genotype at each generation. (C) Wild-type and simr-1 mutant males were raised either at 20°C, a single
generation at 25°C, or following 10 generations of growth at 25°C, and then mated to fog-2 females raised at 20°C.
Brood sizes were scored for 10 fog-2 females, each mated to four males of the indicated genotypes, and demonstrating
that simr-1 male fertility is compromised at 25°C. (D) Wild-type and simr-1 mutant hermaphrodites were raised either
at 20°C, a single generation at 25°C, or following 10 generations of growth at 25°C, and then mated to four pgl-1::gfp
males raised at 20°C. Brood sizes were scored for each of 10 wild-type or simr-1 mutant hermaphrodites, mated to four
pgl-1::gfp males. Only plates with GFP positive progeny were scored. These data indicate that oogenesis of simr-1 is
compromised after multiple generations at 25°C. (E) Number of apoptotic germ cells were counted in a minimum of 20
wild-type and simr-1 mutant gonads using CED-1::GFP engulfment as a marker for apoptotic germ cells. Animals were
raised either at 20°C, or for one, two, four, seven, 10 or 11 generations at 25°C, and imaged approximately 24 hours
after the L4 larval stage. Error bars indicate SEM. n.s. denotes not significant and indicates a p-value > 0.05, * indicates
a p-value ≤ 0.05, *** indicates a p-value ≤ 0.001, **** indicates a p-value ≤ 0.0001. See Supplementary File 8(Manage
et al 2020) for more details regarding statistical analysis.

78

Figure 3–figure supplement 1. hpo-40 does not contribute to the progressive sterility of simr-1
mutants.
Brood sizes of wild-type, hpo-40 single mutants, and simr-1; hpo-40 double mutants were scored for a single generation
at 20°C, followed by 11 generations at 25°C, demonstrating that hpo-40 does not contribute to the progressive sterility
of simr-1 mutants at 25°C. 10 broods were scored for each genotype at each generation.

79

Figure 4. simr-1 mutants have piRNA-related defects.
(A) Images of adult animals, in which the henn-1 mutation weakly desilences the piRNA sensor (top). simr-1[A11V]
(middle) and simr-1[R159C] (bottom) mutants, obtained from an EMS mutagenesis screen of the henn-1; piRNA sensor
strain further desilence the sensor and increase GFP expression. All images were obtained using the same microscope
settings. Scale bars, 50μm. (B) A mating-based approach to reestablish WAGO-class 22G-siRNA production in the
presence and absence of simr-1 and prg-1. Schematic (top) illustrating the three crosses and bar graph (bottom)
showing percentage of fertile and sterile animals from each cross. (C) Live imaging of SIMR-1::GFP (left) and SIMR-
1[R159C]::GFP (right) demonstrate that Tudor domain is critical for SIMR-1 localization to perinuclear foci. Scale bars,
5μm. (D) Brood size was scored for simr-1::gfp and simr-1[R159C]::gfp strains at 20°C, then animals were raised for
11 generations at 25°C. Broods were additionally scored at generations one, five, seven, 10 and 11 at 25°C
demonstrating that the simr-1[R159C]::gfp strain becomes progressively sterile at 25°C, similar to the simr-1 null
mutation, while simr-1::gfp maintains fertility at 25°C similar to wild-type animals. 10 broods were scored for each
genotype at each generation.
80

Figure 4–figure supplement 1. simr-1 mutants do not display defects associated with mutants in the
mutator or ERGO-1 26G-siRNA pathways.
(A) Images of adult animals carrying the piRNA sensor in a wild-type (top), simr-1 mutant (middle), or mut-16 mutant
(bottom) background. The piRNA sensor was maintained in a wild-type background for many generations prior to
introduction of the simr-1 and mut-16 mutations and therefore is likely subject to heritable RNA silencing (RNAe). The
simr-1 mutation is unable to relieve the RNAe-mediated silencing of the wild-type piRNA sensor, in contrast to the simr-
1-mediated desilencing of the henn-1; piRNA sensor (Figure 4A). All images were obtained using the same microscope
settings and processed identically. Scale bars, 50μm. (B) Images of adults carrying the 22G-siR1 sensor, which is
sensitive to perturbations in the ERGO-1 26G-siRNA pathway and the downstream mutator pathway. The simr-1
mutation (middle) was unable to desilence the 22G-siR1 sensor and expresses GFP at levels similar to a wild-type
sensor strain (top). In contrast, mut-16 mutants robustly expressed GFP from the desilenced 22G-siR1 sensor. All
images were obtained using the same microscope settings and processed identically. (C) Animals carrying the simr-1
deletion allele were assayed for their ability to respond to lir-1, hmr-1, or dpy-13 RNAi. “+” indicates an Enhanced RNAi
(Eri) response and “-” indicates a wild-type response. Wild-type and eri-7 deletion animals, which display the Eri
phenotype, are included as controls. (D) Western blot of SIMR-1::GFP and SIMR-1[R159C]::GFP in one-day adult
animals demonstrate that the R159C mutation in the Tudor domain does not affect SIMR-1 protein expression. Actin
protein is shown as a loading control.

A
piRNA sensor
simr-1; piRNA sensor
mut-16; piRNA sensor
B
22G-siR-1 sensor
simr-1; 22G-siR-1 sensor
mut-16; 22G-siR-1 sensor
lir-1 hmr-1 dpy-13
wild-type
eri-7
simr-1
gene
- - -
+ + +
- - -
C
D
135 kDa
46 kDa
-actin
-FLAG
SIMR-1[R159C]::GFP::3xFLAG
SIMR-1::GFP::3xFLAG
81

Figure 5. simr-1 mutants display reduced small RNAs mapping to mutator and piRNA-target genes.
(A) Table indicating the number of genes for which the total small RNA levels are either increased or reduced by at
least two-fold for each indicated mutant. All genes also met the requirements of having at least 10 RPM in either mutant
or control and a DESeq2 adjusted p-value of ≤ 0.05. (B) Box plots displaying total small RNAs levels mapping to genes
from the indicated small RNA pathways in simr-1 mutants compared to wild-type animals raised at 20°C. Details
regarding definition of small RNA target gene classes is provided in the Materials and Methods section. At least 10
RPM in wild-type or simr-1 mutant libraries was required to be included in the analysis. (C) Heat maps displaying total
small RNAs levels targeting mutator-target genes or CSR-1-target genes in simr-1 mutants raised at 20°C, a single
generation at 25°C, or two, seven, or 10 generations at 25°C relative to wild-type at the same temperature and
generation. (D) Venn diagrams indicating overlap of genes depleted of total small RNAs by two-fold or more in mutants
compared to wild-type. (E) Reads per total million reads mapping to piRNA and piRNA-target gene loci in wild-type and
82
simr-1 mutants raised at either 20°C, or for a single generation at 25°C, indicate that piRNAs are not reduced in simr-
1 mutants. Error bars indicate standard deviation of two replicate libraries. (F) Scatter plots display piRNA reads per
million total reads in wild-type and simr-1 mutants (top) and wild-type and prg-1 mutants (bottom). Genes with two-fold
increase in piRNA abundance and DESeq2 adjusted p-value ≤ 0.05 are colored dark red and genes with two-fold
reduction in piRNA abundance and DESeq2 adjusted p-value ≤ 0.05 are colored light blue. The percentage of total
piRNAs with an increase or reduction of greater than two-fold is indicated in the corners of the graph. n.s. denotes not
significant and indicates a p-value > 0.05 and **** indicates a p-value ≤ 0.0001. See Supplementary File 8(Manage et
al 2020) for more details regarding statistical analysis.

83

Figure 5–figure supplement 1. Small RNAs are reduced at many mutator, piRNA, and ERGO-1 target
genes in simr-1 mutants at 25°C.
(A) Box plot displaying total small RNAs levels targeting genes from the indicated small RNA pathways in simr-1
mutants relative to wild-type animals raised for a single generation at 25°C. Details regarding definition of small RNA
target gene classes is provided in the Materials and Methods section. (B) Small RNA distribution across the RDE-1-
target gene Y47H10A.5 in wild-type, simr-1 mutants, prg-1 mutants and mut-16 mutants at 20°C and wild-type, simr-1
mutants, and mut-16 mutants for a single generation at 25°C, shows that small RNAs targeting Y47H10A.5 are
produced independently of simr-1 and prg-1. (C) Venn diagrams indicating overlap of genes depleted of total small
RNAs by two-fold or more in simr-1, prg-1, and mut-16 mutants compared to wild-type. (D) Box plot displaying total
small RNAs levels mapping to piRNA-target genes in simr-1 mutants raised at 20°C or for one, two, seven or 10
generations at 25°C compared to wild-type animals raised at the same temperature for the same number of
generations. piRNA-target genes are defined as those depleted of total small RNAs by two-fold or more in a prg-1
mutant, therefore all piRNA-target genes will have a log2(fold change small RNA abundance) ≤ -1 in a prg-1 mutant
and would fall below the red dotted line.
84

Figure 6. simr-1 mutants display reduced small RNAs mapping to piRNA-dependent transposons
and increased small RNAs mapping to histone genes.
(A) Ratio of transposon-mapping small RNA reads per million total reads in simr-1 mutants and mut-16 mutants raised
at 20°C or a single generation at 25°C compared to wild-type shows that small RNAs mapping to Tc2 and Tc3 depend
on SIMR-1, but those mapping to Tc1 and Tc4v do not. Error bars indicate standard deviation of two replicate libraries.
(B) Venn diagram indicating overlap between genes enriched for small RNAs in simr-1 mutants, prg-1 mutants, and
mut-16 mutants. (C) Venn diagram (top) of the 104 genes enriched for small RNAs in both simr-1 and prg-1 mutants
compared to a list of all histone genes. A pie chart (bottom) of all histone genes shows the number of genes enriched
for small RNAs in prg-1, simr-1, and mut-16 mutants compared to wild-type. (D) Box plot displays small RNAs mapping
to histone genes in simr-1 mutants raised at 20°C, a single generation at 25°C, or two, seven, or 10 generations at
85
25°C, mut-16 mutants at 20°C or 25°C, and prg-1 mutants at 20°C compared to wild-type animals at the same
temperature and generation, demonstrating that small RNAs mapping to histone genes increase at all temperatures in
simr-1 mutants and in prg-1 mutants but not mut-16 mutants. (E) Box plot displays small RNAs mapping to histone
gene classes in simr-1 mutants raised at 20°C or a single generation at 25°C and prg-1 mutants at 20°C compared to
wild-type animals, demonstrating that small RNAs mapping to some histone gene classes increase in both simr-1
mutants and in prg-1 mutants while others increase only in prg-1 mutants. (F) Table indicating the number of genes for
which the mRNA expression is either increased or reduced by at least two-fold for each indicated mutant. All genes
met the requirements of having a DESeq2 adjusted p-value of ≤ 0.05 but no minimum read count was required. (G)
Enrichment analysis (log2(fold enrichment)) examining the overlap of genes up and down-regulated in simr-1 mutants
with known targets of the CSR-1, mutator, PRG-1 and ALG-3/4 small RNA pathways and oogenesis and
spermatogenesis-enriched genes. Color of boxes correlates with fold enrichment (red) or depletion (blue). Statistical
significance for enrichment was calculated using the Fisher’s Exact Test function in R. (H) Box plot displays mRNA
expression in simr-1 (blue) or mut-16 (grey) relative to wild-type for genes that are enriched or depleted of small RNAs
in the same mutants. (I) Box plot displays histone mRNA expression in simr-1 (blue) or mut-16 (grey) relative to wild-
type, demonstrating that histone mRNA expression is reduced in simr-1 mutant animals. n.s. denotes not significant
and indicates a p-value > 0.05, * indicates a p-value ≤ 0.05, ** indicates a p-value ≤ 0.01, *** indicates a p-value ≤
0.001, **** indicates a p-value ≤ 0.0001. See Supplementary File 8(Manage et al 2020) for more details regarding
statistical analysis.

86

Figure 6–figure supplement 1. Small RNAs mapping to piRNA target transposons are reduced and
small RNAs mapping to histone genes are increased in simr-1 mutants.
(A) Heat maps displaying small RNAs levels targeting mutator-dependent transposons and repeats in prg-1 and simr-
1 mutants raised at 20°C, and simr-1 mutants raised at 25°C for one, two, seven, or 10 generations compared to wild-
type at the same temperature and generation. (B) Venn diagrams showing the overlap of mutator-dependent
transposons (left) or mutator-dependent repeats (right) depleted of small RNAs in simr-1 mutants compared to those
depleted of small RNAs in prg-1 mutants at 20°C. (C,D) Small RNA distribution across two transposon loci in wild-type,
simr-1 mutants, mut-16 mutants, and prg-1 mutants at 20°C and in wild-type, simr-1 mutants, and mut-16 mutants after
a single generation at 25°C. The Tc2 locus (C) requires SIMR-1, PRG-1 and MUT-16 for wild-type small RNA levels,
while the Tc1 locus (D) requires MUT-16 but not SIMR-1 or PRG-1 for small RNA production. (E) Small RNA distribution
across a histone gene cluster in wild-type, simr-1 mutants, mut-16 mutants, and prg-1 mutants at 20°C and in wild-
type, simr-1 mutants, and mut-16 mutants after a single generation at 25°C, shows that siRNAs targeting histone genes
87
increase in simr-1 mutants and prg-1 mutants but not mut-16 mutants. (F) Venn diagram showing the overlap of genes
with increased mRNA expression compared to genes with reduced small RNAs in simr-1 mutants raised at 20°C. (G)
Box plot displaying mRNAs mapping to the histone gene classes in simr-1 mutants raised at 20°C, shows that distinct
histone gene classes have reduced mRNA expression in simr-1 mutants.

88

Figure 7. SIMR-1 localizes to foci adjacent to P granules and Mutator foci.
(A) Immunostaining of SIMR-1 (green) with MUT-16 (red, top left), PGL-1 (red, top right), ZNFX-1 (red, bottom left),
and RSD-2 (red, bottom right) demonstrates that SIMR-1 localizes to foci near Mutator foci (MUT-16), P granules (PGL-
1), and Z granules (ZNFX-1) but overlaps most substantially with RSD-2 foci. Arrow indicates an example of a single Z
granule associated with two SIMR-1 foci. (B) Bar graph showing distance between the centers of fluorescence for
indicated proteins to SIMR-1 (mean +/- SEM). See Materials and Methods for description of quantification methods.
n.s. denotes not significant and indicates a p-value > 0.05, * indicates a p-value ≤ 0.05, ** indicates a p-value ≤ 0.01,
**** indicates a p-value ≤ 0.0001. See Supplementary File 8(Manage et al 2020) for more details regarding statistical
analysis. (C) Immunostaining of SIMR-1 (green), ZNFX-1 (red), and PGL-1 (white) allows for visualization of the stacked
SIMR/Z granule/P granule foci. All images are projections of 3D images following deconvolution. DAPI is blue in all
panels and scale bars are 5μm.

A
SIMR-1
RSD-2
B
SIMR-1
ZNFX-1
SIMR-1
PGL-1
SIMR-1
MUT-16
SIMR-1
ZNFX-1
PGL-1
C
0
0.05
0.1
0.15
0.2
0.25
0.3
RSD-2 MUT-16 ZNFX-1 PGL-1
n.s.
Distance to SIMR-1 foci (nm)
**
*
****
*
89

Figure 7–figure supplement 1. SIMR-1 and PRG-1 localize independently and to distinct granules.
(A) Immunostaining of SIMR-1::GFP (green) and PRG-1::mKate (red) demonstrates that SIMR-1 and PRG-1 localize
near one another but their localization is not coincident. (B) Live imaging of SIMR-1::GFP in a prg-1 mutant (top) and
PRG-1::mKate in a simr-1 mutant (bottom) indicate that prg-1 is not required for SIMR-1 foci formation, nor is simr-1
required for PRG-1 localization to P granules. (C) Western blot of mKate2::3xMyc::PRG-1 in a wild-type or simr-1
mutant background demonstrate that the simr-1 mutation does not affect PRG-1 protein expression. Actin protein is
shown as a loading control. Scale bars, 5μm.

simr-1 mKate2::PRG-1
A
C
SIMR-1::GFP prg-1
B
135 kDa
46 kDa
-actin
-Myc
mKate2::3xMyc::PRG-1
simr-1 wild-type
SIMR-1
PRG-1
90
Chapter III
Proximity labeling to identify SIMR-1 protein interactors

Introduction
SIMR-1 (siRNA-defective and mortal germline) was first identified in C. elegans
through proteomic analysis of MUT-16, a critical component in siRNA amplification.
SIMR-1 was a completely uncharacterized protein, and unlike most other RNA silencing
pathway mutants, simr-1 is not RNAi defective. However, as is characteristic of mutations
in the mutator pathway, simr-1 animals express a transgenerational sterility phenotype at
elevated temperatures and become sterile within 11 generations (Manage et al., 2020;
Ketting, 1999; Phillips, 2012; Zhang, 2011). Loss of SIMR-1 does not have any effect on
piRNA production or PRG-1 expression but does lead to an inability to produce the large
numbers of siRNAs from piRNA-target loci necessary for effective, robust silencing. This
suggests that SIMR-1 is required at a step between PRG-1 targeting and biogenesis of
siRNAs by the mutator complex (Manage et al., 2020). SIMR-1, along with a small RNA
factor required for exogenous RNAi, RSD-2 (Han, 2008; Sakaguchi, 2014), were found
to localize to distinct perinuclear foci in the C. elegans germline called SIMR foci (Manage
et al., 2020). SIMR foci localize adjacent to, but distinct from other known granules
(Mutator foci, P granule and Z granules) as part of C. elegans nuage (Manage et al.,
2020). Currently, SIMR-1 and RSD-2 are the only known components of SIMR foci.
There are several methods that can be used to identify protein-protein interactions,
but isolation of those interactions by methods such as affinity purification and yeast two-
hybrids, must be conducted in vitro or under non-physiological conditions (Kim and Roux,
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2016; Kim et al., 2016). Enzyme-catalyzed proximity labeling is an alternate form of
proteomic analysis that addresses some of the deficiencies of these other methods and
is constantly being improved and adapted upon as it is utilized by the scientific community
(Branon et al., 2018). In addition to being effective in vivo, proximity labeling can also
detect protein interactions that have low affinities, are transient, and can also identify
protein constituents of subcellular domains (Kim and Roux, 2016; Kim et al., 2016).
Proximity labeling uses a promiscuous enzyme fused to a protein of interest, that
when supplemented with a small molecule substrate such as biotin, catalyzes a covalent
labeling reaction that tags proteins within proximity of the fusion protein (Branon et al.,
2018; Hung et al., 2016; Lam et al., 2015; Sears et al., 2019; V’kovski et al., 2020). These
biotinylated proteins can then be isolated by affinity purification and identified via mass
spectrometry (mass spec). The high affinity of streptavidin and avidin for biotinylated
proteins allows for fast, specific isolation of tagged proteins and because this is a covalent
modification, biotinylated proteins can withstand stringent lysis and wash conditions (Kim
and Roux, 2016). There are several promiscuous enzymes available for use in proximity
labeling such as the soybean derived ascorbate peroxidase APEX2 and the promiscuous
E. coli biotin ligase BioID, but each has their own benefits and limitations. The main
advantage to APEX2 is speed, with tagging occurring within 1 minute of system activation.
However, APEX2 requires the use of H2O2 which is toxic to most model organisms
(Branon et al., 2018; Kim et al., 2016) In contrast, BioID is non-toxic, using biotin as its
labeling reagent, and has been used in many models but requires long periods of labeling,
anywhere from 18-24 hours (Branon et al., 2018; Roux et al., 2012). TurboID is a relatively
new proximity labeling system that improves upon both APEX2 and BioID. TurboID was
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generated by directed evolution of the E. coli biotin ligase BirA (also used to generate
BioID) and allows for the usage of non-toxic biotin as an enzyme substrate with a much
shorter labeling time of 10 minutes (Branon et al., 2018). Proximity-dependent
biotinylation occurs by the ligase catalyzing the formation of biotin-5’-AMP anhydride,
which diffuses from the active site to biotinylate nearby proteins on nucleophilic residues
such as lysine (Branon et al., 2018).
Though streptavidin pull downs of biotinylated proteins are efficient and fast,
results from mass spec can be convoluted by endogenously biotinylated proteins. To
reduce the level of endogenously biotinylated proteins, I made use of the auxin-inducible
degradation (AID) system to target the C. elegans biotin ligase, BPL-1, for degradation.
The AID system consists of heterologously expressed Arabidopsis thaliana TIR1 that in
the presence of auxin (indole-3-acetic acid) interacts with endogenous components to
form a functional E3 ubiquitin ligase. This leads to polyubiquitination of any degron tagged
protein targets, and subsequent degradation by the proteosome (Zhang et al., 2015). Use
of this system in C. elegans allows for rapid, reversible, auxin-dependent degradation of
degron tagged proteins, in this case endogenous biotin ligases.
Here, I developed TurboID for use in proximity labeling of SIMR-1 protein
interactors in the C. elegans germline, in conjunction with the AID system targeting the
endogenous biotin ligase, BPL-1. These two methodologies in tandem will allow for
identification of protein-protein interactions between SIMR-1 and unknown members of
SIMR foci, as well as interactors from neighboring germ granules, while reducing the level
of endogenously biotinylated proteins.

93
Results
Fusion of a proximity labeling enzyme to SIMR-1
In an effort to further identify and characterize SIMR foci components as well as
identify direct interactors of SIMR-1, I conducted an immunoprecipitation mass spec
experiment using epitope tagged versions of SIMR-1. However, perhaps due to the
transient or weak interactions of SIMR-1 with other proteins in the perinuclear space,
these experiments were unsuccessful. In order to utilize mass spec, I adapted proximity
labeling (Figure 1) for use in the C. elegans germline and used CRISPR to endogenously
tag SIMR-1 with 3xHA and the TurboID biotin ligase.
To ensure that fusing SIMR-1 with HA-TurboID did not alter the perinuclear
localization, I performed immunofluorescence. This experiment confirmed that SIMR-
1::3xHA::TurboID localization mirrors that of SIMR-1::mCherry (Manage et al., 2020) and
is found at punctate foci surrounding germline nuclei (Figure 3). Next, I sought to
determine whether SIMR-1::3xHA::TurboID could biotinylate nearby proteins. To this end,
I used the biotin-binding protein, Streptavidin, covalently attached to a fluorescent label,
to determine the location of biotin-labeled proteins in the germline of SIMR-
1::3xHA::TurboID animals. I found a strong signal for biotinylated proteins in proximity of
SIMR-1 (Figure 3). This data suggests that the SIMR-1::3xHA::TurboID fusion protein is
capable of biotinylating nearby proteins or itself and that fusion of TurboID to SIMR-1 did
not disrupt its normal expression in the perinuclear space.

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TurboID biotinylates nearby proteins
In order to further confirm biotinylation activity, and to compare the presence or
absence of biotinylated proteins in wild-type versus simr-1::3xHA::TurboID animals, I
conducted streptavidin chemiluminescent blotting. The animals were raised for 2
generations on biotin-auxotrophic E. coli (MG1655bioB:kan) to deplete the animals of
excess biotin, instead of the usual biotin-producing E. coli strain, OP50 (Figure
4A)(Sulston et al., 1983). The results of the blot indicate that there are biotinylated
proteins present in the simr-1::3xHA::TurboID that are not present in wild-type animals,
including SIMR-1 and at least one unknown protein (Figure 4B and 4C). I also confirmed
the identity of SIMR-1 by Western blotting with an anti-HA horseradish peroxidase
antibody (Figure 4D). There may be additional biotinylated proteins in the simr-
1::3xHA::TurboID strain that are expressed below the level of detection in this streptavidin
blot. However, there are also several prominent biotinylated proteins that are present in
both wild-type and simr-1::3xHA::TurboID animals. In previous work, it has been
demonstrated that BPL-1, a C. elegans ortholog of mammalian holocarboxylase
synthetase involved in fatty acid biosynthesis, can biotinylate multiple endogenous
proteins, including pod-2 (acetyl-CoA carboxylase), pyc-1 (pyruvate carboxylase), pcca-
1 (propionyl-CoA carboxylase subunit A) and mccc-1 (3-methylcrotonyl-CoA carboxylase)
(Figure 4B)(Watts et al., 2018). Based on their predicted sizes, I can identify enzymes
PYC-1, PCCA-1 and MCCC-1 in the streptavidin blot (Watts et al., 2018). This data
together suggests that simr-1::3xHA::TurboID animals biotinylate additional proteins, not
seen in wild-type animals.

95
BPL-1 degradation by the AID system
Because the majority of biotinylated proteins in the simr-1::3xHA::TurboID strain
are proteins biotinylated by the BPL-1 biotin ligase rather than the SIMR-
1::3xHA::TurboID, I sought to reduce the expression of BPL-1 and correspondingly
reduce the biotinylation of its target proteins. While there are a number of methods that
could be used to reduce BPL-1 expression, I chose the AID system for the following
reasons – 1) Many methods for protein depletion are limiting, including RNAi or gene
disruption as they may disrupt proteins or pathways necessary for animal development
or survival (Cheng et al., 2013; Shen et al., 2014); 2) Use of an auxin-inducible system
allows for precise destruction of degron tagged proteins and allows for a great deal of
temporal and spatial control through expression by specific promoters and introduction of
auxin at various timepoints (Ashley et al., 2021; Yesbolatova et al., 2020; Zhang et al.,
2015).
To generate degron-tagged BPL-1, I used CRISPR to fuse a degron tag to BPL-1
at the endogenous locus in the simr-1::3xHA::TurboID strain. I then generated two strains
where I introduced a heterologously expressed Arabidopsis thaliana TIR1 into simr-
1::3xHA::TurboID, one under a germline promoter and another under a pan-somatic
promoter, bringing together all the components necessary for directed degradation of
BPL-1 (Figure 2). I began testing of the AID system using the animals expressing pan-
somatic TIR1, simr-1::3xHA::TurboID; bpl-1::deg; Peft-3::TIR1::mRuby, which I exposed
to 100mM auxin and following a two hour exposure time, collected the animals for
streptavidin blotting. There appeared to be very little, if any, change in expression of
endogenously biotinylated proteins (Figure 4C). There are many possible explanations
96
for this lack of depletion, the most likely being that though I activated the AID system for
two hours, that was not enough time for degradation of previously biotinylated
endogenous proteins, at least to a level that would make a significant difference. As we
are only knocking down the biotinylase instead of the biotinylated proteins themselves,
degradation of the endogenously biotinylated proteins may take longer than the amount
of time I subjected the animals to auxin. It is also possible that use of the strain expressing
pan-somatic TIR1 was insufficient to knockdown BPL-1and may be more effective with
TIR1 expression throughout both somatic and germline tissue.

Yeast two-hybrid results for SIMR-1
As another way to identify SIMR-1 interactors to validate the results of our
proximity labeling, we conducted a yeast two-hybrid. Three high-confidence protein
interactors of SIMR-1 were found - F45D3.4, HRDE-2 and MBK-2 (Supplementary file 1).
F45D3.4 is an uncharacterized protein with no discernible domains or protein family
membership. Preliminary work ongoing in the lab has demonstrated that deletion of
F45D3.4 leads to a marked decrease in fertility at permissive temperatures and, as with
simr-1 mutant animals, we observe a drastic decrease in fertility when exposed to
elevated temperatures of 25°C (Manage et al., 2020). Additionally, like the simr-1 mutant,
preliminary data has shown that the F45D3.4 deletion mutant responds like wild-type to
both germline and somatic RNAi. HRDE-2 is a known small RNA factor that was
previously identified in a genetic screen for RNAi inheritance (Spracklin et al., 2017). hrde-
2 mutant animals were found to have temperature sensitive, mortal germline phenotype,
which is characteristic of RNAi inheritance factor mutants (Spracklin et al., 2017). HRDE-
97
2 was also found to localize to distinct, perinuclear foci in the C. elegans germline, with
some preliminary data from our lab indicating it localizes to SIMR foci (Spracklin et al.,
2017). MBK-2 is a dual-specificity YAK-1-related(DYRK) kinase that has been shown to
directly phosphorylate oocyte proteins for degradation as oocytes transition into
embryogenesis as well as phosphorylate P granule components MEG-1 and MEG-3,
promoting P granule disassembly (Stitzel 2006,Shirayama 2006,Wang 2014).
Established data as well as preliminary data collected by our lab suggests that F45D3.4,
HRDE-2 and MBK-2 are strong candidates for interactors of SIMR-1 based on their
phenotypes or involvement with P granules and the piRNA pathway. Further
characterization of each candidate as well as validation of yeast two-hybrid results by
proximity labeling and mass spec of SIMR-1 interactors will be necessary.

Discussion
In order to ensure high enrichment of biotinylated proteins during streptavidin pull
down as well as reducing levels of endogenously biotinylated proteins, there are many
adjustments that can be made to this methodology. As with previous studies, simr-
1::3xHA::TurboID; bpl-1::deg; Peft-3:TIR1::mRuby animals were grown on biotin
auxotrophic bacteria for two generations to deplete them of excess biotin. However, a
recent study has suggested that the addition of exogenous biotin may increase protein
biotinylation (Artan et al., 2021). Various concentrations and time points were tested for
addition of supplemental biotin to worms grown on biotin-auxotrophic bacteria and it was
found that 1mM biotin supplemented two hours prior to animal collection led to robust
biotinylation (Artan et al., 2021). However, these adjustments may also lead to excess
98
free biotin that will need to be filtered out before exposure of animal lysates to streptavidin-
covered beads. Supplemented free biotin will likely compete with biotinylated proteins for
available streptavidin binding sites. Another alternative may be to utilize commercially
available high-capacity streptavidin bound beads which will likely ensure isolation of
biotinylated proteins, though with potentially higher background.
To further reduce endogenously biotinylated proteins, optimization of the AID
system will be necessary. Varying the auxin concentrations and exposure times may
provide the necessary stringency when reducing endogenously biotinylated proteins.
Though I exposed the simr-1::3xHA::TurboID; bpl-1::deg; Peft-3::TIR1::mRuby animals
to auxin two hours prior to collection, that may not have been enough time to see
decreases in endogenous biotinylated protein expression. Identifying the ideal AID
system activation time will be critical to ensuring that background signal will be reduced
as much as possible during mass spec. Additionally, there has been recent work further
optimizing the AID system as well as the creation of complementary tools to enable better
targeted degradation as well as improved imaging-based measures of target depletion
(Ashley et al., 2021). The development of degron tags bound to alternative fluorescent
proteins such as blue fluorescent protein (BFP) allow for the depletion phenotype to be
confirmed through degradation of the BFP tag, without interfering with both red and green
channels typically used in experiments (Ashley et al., 2021).
Another major avenue to pursue in boosting enrichment and reducing background
would be to make modifications to the procedures following animal collection, during
affinity purification with streptavidin-coated beads. Increasing the number of streptavidin-
coated beads used, as well as ensuring wash steps are more stringent will likely lead to
99
more promising results in the future. As optimization of this protocol is still in the relatively
early stages, there is still a great deal of work to be done regarding isolation and
identification of SIMR-1 interactors via TurboID. Ideally, upon completion of this
methodology and the return of reliable, replicable results, this combination TurboID and
AID system can be utilized to identify interactors of many proteins found in germ granules
and the perinuclear space.

Materials and Methods
Strains
The wild-type C. elegans strain used was N2. Worms were raised at 20°C under
standard conditions (Brenner, 1974). All strains used for this project are listed in
Supplementary file 2.

Plasmid and strain construction
Plasmid containing 3xHA::TurboID was purchased from Addgene (# 118220).
simr-1::3xHA::TurboID was generated by CRISPR injection of melted PCR donor repair
template and RNA guide into wild-type animals according to published protocols
(Dickinson 2013,Dickinson 2015,Dokshin 2018,Paix 2015,Paix 2017,Ward 2015,Ghanta
2020,Dokshin 2018,Ghanta 2020). CRISPR injection mix included 25ng/µl melted PCR
repair oligo, .04ng/µl simr-1 guide RNA, 15ng/µl co-injection guide oligo, 110ng/µl co-
injection repair oligo (Supplementary file 3). simr-1::3xHA::TurboID; bpl-1::deg was
generated by CRISPR injection that included 110ng/µl bpl-1:deg repair oligo, .04ng/µl bpl-
100
1 guide RNA, 15ng/µl co-injection guide oligo, 110ng/µl co-injection repair oligo
(Supplementary file 3).

Antibody staining and imaging
Immunofluorescence was conducted on one-day old adults (~24 hours after L4),
dissected in egg buffer containing 0.1% Tween-20 and fixed in 1% formaldehyde in egg
buffer as described (Phillips et al., 2009). Dissected gonads were immunostained with
mouse anti-HA (3F10, Sigma 11867423001) and streptavidin (AlexaFluor 488,
ThermoFisher S11223). Alexa -Fluor secondary antibodies were purchased and utilized
from ThermoFisher Scientific. Imaging was conducted on DeltaVision Elite microscope
(GE Healthcare) using 60x N.A. 1.42 oil-immersion objective. Image analysis was
performed using SoftWoRx package and when data stacks were collected, deconvolution
was conducted and images were presented as max intensity projections, followed by
pseudo-coloring using Adobe Photoshop.

Yeast two-hybrid
Yeast two-hybrid was conducted by Hybrigenics services.

Streptavidin bead purification of biotinylated protein protocol
Materials and Reagents
1. Indole-3-acetic acid (auxin)(GoldBio, cat. No. I-110-100)
2. Dynabeads M-280 Streptavidin beads (ThermoFisher, cat. No. 11205D)
3. Bovine serum albumin (VWR, cat. No. 97061-422)
101
4. Pierce proteinase inhibitor tablet (ThermoFisher, cat. No. A32965)

Procedure
1. Grow synchronized animals for two generations on biotin-auxotrophic E. coli
(MG1655bioB:kan).
Day before collection:
2. Wash mortar and pestle in hot water, 10% SDS, hot water, ethanol, let air dry.
3. Label tubes and place in cold room: lysate, supernatant, input, and final for
each sample .
Day of purification:
4. Collect animals as day 1 adults (68 hours post L1 plating).
• 2 hours before collection, dose animals with 100mM auxin.
5. Prep 50mL lysis buffer with protease (1 proteinase inhibitor tablet, crush and
dissolve) KEEP COLD.
6. Prep worms (during 1hr rotation step for beads).
• Cool grinders with liquid nitrogen.
• Take worm pellets directly from -80°C and gently crush in mortar.
• Grind to powder and add 1.5mL of lysis buffer with protease inhibitor.
• Once slushy, cover with saran wrap and move to 4°C.
7. IN COLD ROOM: Add all homogenate to the corresponding two “lysate” tubes,
centrifuge for 10min at max speed.
8. Prep beads: Dynabeads M-280 Streptavidin (during 10-minute centrifugation,
can be done at room temp).
102
• Transfer desired volume of beads to a tube.
§ 40µL (.2mg beads) bead slurry per sample with extra for error
margin. Mark slurry level before next step.
• Add an equal volume of washing buffer (1x PBS, pH 7.4) or at least 1mL
and mix (vortex for 5 sec, or keep on a roller for at least 5 minutes).
• Place the tube on a magnet rack for 1 min and discard the supernatant.
• Remove the tube from the magnet and resuspend the washed beads in the
same volume of 1x PBS as the initial volume of beads taken from the vial.
9. Save 40µl of the supernatant in INPUT tubes.
10. Transfer rest of supernatant to “supernatant” tubes and add 40µl bead slurry.
• Avoid fat layer when transferring supernatant.
11. ROTATE 30 minutes at RT (1 hour at 4°C is ok).
12. Separate the protein-coated beads with a magnet for 2-3 minutes.
13. Wash the coated beads 4 times in 1ml PBS containing 0.1% BSA (vortex 5 sec
or roll for at least 5 minutes).
14. Resuspend to the desired concentration for your application.
15. Submit streptavidin beads directly to mass spec facility.

References
Artan, M., Barratt, S., Flynn, S.M., Begum, F., Skehel, M., Nicolas, A., and Bono, M. de
(2021). Interactome analysis of C. elegans synapses by TurboID-based proximity
labeling. Biorxiv 2021.04.01.438103.

103
Ashley, G.E., Duong, T., Levenson, M.T., Martinez, M.A.Q., Johnson, L.C., Hibshman,
J.D., Saeger, H.N., Palmisano, N.J., Doonan, R., Martinez-Mendez, R., et al. (2021). An
expanded auxin-inducible degron toolkit for Caenorhabditis elegans. Genetics 217,
iyab006.
Branon, T.C., Bosch, J.A., Sanchez, A.D., Udeshi, N.D., Svinkina, T., Carr, S.A.,
Feldman, J.L., Perrimon, N., and Ting, A.Y. (2018). Efficient proximity labeling in living
cells and organisms with TurboID. Nature Biotechnology 36, 880–887.
Brenner, S. (1974). THE GENETICS OF CAENORHABDITIS ELEGANS. Genetics 77,
71–94.
Cheng, Z., Yi, P., Wang, X., Chai, Y., Feng, G., Yang, Y., Liang, X., Zhu, Z., Li, W., and
Ou, G. (2013). Conditional targeted genome editing using somatically expressed TALENs
in C. elegans. Nat Biotechnol 31, 934–937.
Dickinson, D.J., Ward, J.D., Reiner, D.J., and Goldstein, B. (2013). Engineering the
Caenorhabditis elegans genome using Cas9-triggered homologous recombination. Nat
Methods 10, 1028–1034.
Dickinson, D.J., Pani, A.M., Heppert, J.K., Higgins, C.D., and Goldstein, B. (2015).
Streamlined Genome Engineering with a Self-Excising Drug Selection Cassette. Genetics
200, 1035–1049.
Dokshin, G.A., Ghanta, K.S., Piscopo, K.M., and Mello, C.C. (2018a). Robust Genome
Editing with Short Single-Stranded and Long, Partially Single-Stranded DNA Donors in
Caenorhabditis elegans. Genetics 210, 781–787.
Dokshin, G.A., Ghanta, K.S., Piscopo, K.M., and Mello, C.C. (2018b). Robust genome
editing with short single-stranded and long, partially single-stranded DNA donors in C.
elegans. Biorxiv 352260.
Fields, S., and Song, O. (1989). A novel genetic system to detect protein–protein
interactions. Nature 340, 245–246.
Ghanta, K.S., and Mello, C.C. (2020a). Melting dsDNA Donor Molecules Greatly
Improves Precision Genome Editing in Caenorhabditis elegans. Genetics 216, 643–650.
Ghanta, K.S., and Mello, C.C. (2020b). Melting dsDNA donor molecules potentiates
precision genome editing in C. elegans. BioRxiv 1–35.
Han, W., Sundaram, P., Kenjale, H., Grantham, J., and Timmons, L. (2008). The
Caenorhabditis elegans rsd-2 and rsd-6 Genes Are Required for Chromosome Functions
During Exposure to Unfavorable Environments. Genetics 178, 1875–1893.
104
Hung, V., Udeshi, N.D., Lam, S.S., Loh, K.H., Cox, K.J., Pedram, K., Carr, S.A., and Ting,
A.Y. (2016). Spatially resolved proteomic mapping in living cells with the engineered
peroxidase APEX2. Nature Protocols 11, 456–475.
Kim, D.I., and Roux, K.J. (2016). Filling the Void: Proximity-Based Labeling of Proteins in
Living Cells. Trends in Cell Biology 26, 804–817.
Kim, D.I., Jensen, S.C., Noble, K.A., Kc, B., Roux, K.H., Motamedchaboki, K., and Roux,
K.J. (2016). An improved smaller biotin ligase for BioID proximity labeling. Molecular
Biology of the Cell 27, 1188–1196.
Lam, S.S., Martell, J.D., Kamer, K.J., Deerinck, T.J., Ellisman, M.H., Mootha, V.K., and
Ting, A.Y. (2015). Directed evolution of APEX2 for electron microscopy and proximity
labeling. Nature Methods 12, 51–54.
Manage, K.I., Rogers, A.K., Wallis, D.C., Uebel, C.J., Anderson, D.C., Nguyen, D.A.H.,
Arca, K., Brown, K.C., Rodrigues, R.J.C., Albuquerque, B.F.M. de, et al. (2020). A Tudor
domain protein, SIMR-1, promotes siRNA production at piRNA-targeted mRNAs in C.
elegans. ELife 9, 1–33.
Paix, A. (2015). High Efficiency, Homology-Directed Genome Editing in Caenorhabditis
elegans Using CRISPR-Cas9 Ribonucleoprotein Complexes. Genetics Investigation 1–
26.
Paix, A., Folkmann, A., and Seydoux, G. (2017). Precision genome editing using
CRISPR-Cas9 and linear repair templates in C. elegans. Methods 121–122, 86–93.
Phillips, C.M., McDonald, K.L., and Dernburg, A.F. (2009). Cytological Analysis of Meiosis
in Caenorhabditis elegans. pp. 171–195.
Phillips, C.M., Montgomery, T.A., Breen, P.C., and Ruvkun, G. (2012). MUT-16 promotes
formation of perinuclear mutator foci required for RNA silencing in the C. elegans
germline. Genes & Development 26, 1433–1444.
Roux, K.J., Kim, D.I., Raida, M., and Burke, B. (2012). A promiscuous biotin ligase fusion
protein identifies proximal and interacting proteins in mammalian cells. The Journal of
Cell Biology 196, 801–810.
Sakaguchi, A., Sarkies, P., Simon, M., Doebley, A.L., Goldstein, L.D., Hedges, A.,
Ikegami, K., Alvares, S.M., Yang, L., LaRocque, J.R., et al. (2014). Caenorhabditis
elegans RSD-2 and RSD-6 promote germ cell immortality by maintaining small interfering
RNA populations. 111, E4323–E4331.
Sears, R.M., May, D.G., and Roux, K.J. (2019). BioID as a Tool for Protein-Proximity
Labeling in Living Cells. (Humana Press), pp. 299–313.
105
Shen, Z., Zhang, X., Chai, Y., Zhu, Z., Yi, P., Feng, G., Li, W., and Ou, G. (2014).
Conditional Knockouts Generated by Engineered CRISPR-Cas9 Endonuclease Reveal
the Roles of Coronin in C. elegans Neural Development. Dev Cell 30, 625–636.
Shirayama, M., Soto, M.C., Ishidate, T., Kim, S., Nakamura, K., Bei, Y., Heuvel, S. van
den, and Mello, C.C. (2006). The Conserved Kinases CDK-1, GSK-3, KIN-19, and MBK-
2 Promote OMA-1 Destruction to Regulate the Oocyte-to-Embryo Transition in C.
elegans. Curr Biol 16, 47–55.
Spracklin, G., Fields, B., Wan, G., Becker, D., Wallig, A., Shukla, A., and Kennedy, S.
(2017). The RNAi Inheritance Machinery of Caenorhabditis elegans. Genetics 206, 1403–
1416.
Stitzel, M.L., Pellettieri, J., and Seydoux, G. (2006). The C. elegans DYRK Kinase MBK-
2 Marks Oocyte Proteins for Degradation in Response to Meiotic Maturation. Curr Biol
16, 56–62.
Sulston, J.E., Schierenberg, E., White, J.G., and Thomson, J.N. (1983). The embryonic
cell lineage of the nematode Caenorhabditis elegans. Dev Biol 100, 64–119.
Sundby, A.E., Molnar, R.I., and Claycomb, J.M. (2021). Connecting the Dots: Linking
Caenorhabditis elegans Small RNA Pathways and Germ Granules. Trends Cell Biol 31,
387–401.
Tijsterman, M., May, R.C., Simmer, F., Okihara, K.L., and Plasterk, R.H.A. (2004). Genes
Required for Systemic RNA Interference in Caenorhabditis elegans. Curr Biol 14, 111–
116.
Uebel, C.J., Anderson, D.C., Mandarino, L.M., Manage, K.I., Aynaszyan, S., and Phillips,
C.M. (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 Genetics 14, e1007542-22.
V’kovski, P., Steiner, S., and Thiel, V. (2020). Proximity Labeling for the Identification of
Coronavirus-Host Protein Interactions. Methods in Molecular Biology (Clifton, N.J.) 2203,
187–204.
Wan, G., Fields, B.D., Spracklin, G., Shukla, A., Phillips, C.M., and Kennedy, S. (2018).
Spatiotemporal regulation of liquid-like condensates in epigenetic inheritance. Nature 1–
23.
Ward, J.D. (2015). Rapid and Precise Engineering of the Caenorhabditis elegans
Genome with Lethal Mutation Co-Conversion and Inactivation of NHEJ Repair. Genetics
199, 363–377.
106
Watts, J.S., Morton, D.G., Kemphues, K.J., and Watts, J.L. (2018). The biotin-ligating
protein BPL-1 is critical for lipid biosynthesis and polarization of the Caenorhabditis
elegansembryo. The Journal of Biological Chemistry 293, 610–622.
Yesbolatova, A., Saito, Y., Kitamoto, N., Makino-Itou, H., Ajima, R., Nakano, R., Nakaoka,
H., Fukui, K., Gamo, K., Tominari, Y., et al. (2020). The auxin-inducible degron 2
technology provides sharp degradation control in yeast, mammalian cells, and mice.
Nature Communications 1–13.
Zhang, C., Montgomery, T.A., Fischer, S.E.J., Garcia, S.M.D.A., Riedel, C.G., Fahlgren,
N., Sullivan, C.M., Carrington, J.C., and Ruvkun, G. (2012). The Caenorhabditis elegans
RDE-10/RDE-11 Complex Regulates RNAi by Promoting Secondary siRNA
Amplification. Current Biology 22, 881–890.
Zhang, L., Ward, J.D., Cheng, Z., and Development, A.D. (2015). The auxin-inducible
degradation (AID) system enables versatile conditional protein depletion in C. elegans.
Dev.Biologists.Org.

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Figures and Figure Legends

Figure 1. Proximity labeling with TurboID identifies protein-protein interactors by biotinylation.
The promiscuous biotin-ligase TurboID, when bound to a protein of interest, biotinylates nearby interactors that can
then be identified by mass spectrometry following isolation of biotinylated proteins.

10 20 30 40 50 60 70 80 90 100
Relative Intensity
m/z
Protein of interest -
Biotin -
Substrate biotin is utilized
in proximity-dependent
biotinylation of nearby
proteins
Isolation of
denatured
biotinylated
proteins with
streptavidin beads
TurboID fused to
protein of interest
by CRISPR
TurboID Biotin Ligase-
Trypsin digestion of
isolated proteins and
MS identification
108

Figure 2. The AID system allows for fast, and reversible knockdown of degron tagged proteins.
In the presence of auxin (indole-3-acetic acid), the heterologously expressed TIR1 forms a complex with endogenous
proteins to create a functional E3 ubiquitin ligase. Any protein tagged with a degron sequence will be ubiquinated by
the E3 complex for destruction by the proteosome.

TIR1
BPL-1
Ub
Ub
Ub
Degron
BPL-1
Auxin
TIR1
109

Figure 3. SIMR-1 localization is unchanged following fusion to the TurboID biotin ligase.
Immunostaining of SIMR-1(red) demonstrates that SIMR-1 continues to localize to perinuclear foci following fusion to
TurboID. Biotinylation of proteins surrounding SIMR-1 foci (green) indicates TurboID biotinylation is active. All images
are projections of 3D images following deconvolution. DAPI is blue. Scale bars, 5µm.

DAPI Biotin SIMR-1::3xHA::TurboID Merge
110

Figure 4. SIMR-1 bound to TurboID leads to biotinylation of proteins in addition to those
endogenously biotinylated.
(A) Schema for worms grown for two generations on biotin-auxotrophic bacteria before collection for streptavidin blotting
during protocol optimization. (B) simr-1::3xHA::TurboID animals contain additional biotinylated proteins not found in
wild-type animals. (C) Knockdown of BPL-1 by the AID system with 100mM auxin for two hours does not show
significant changes in expression of endogenously biotinylated proteins. (D) Anti-HA Horse Radish Peroxidase Western
blot targeting SIMR-1::3xHA::TurboID.

111
Supplementary file 1. Yeast two-hybrid
results
Gene Name
Number of Y2H
Interactions
C. elegans - F45D3.4 6
C. elegans - hrde-2 143
C. elegans - mbk-2 2

Supplementary file 1. Yeast two-hybrid of SIMR-1 identified three potential interactors.

112

Supplementary file 2. Strains used for this study.
Strain Designation Description
N2 wild-type
CA1199 ieSi38[Psun-1::TIR1::mRuby::sun-1 3' UTR] IV
CA1200 ieSi57[Peft-3::TIR1::mRuby::unc-54 3' UTR] II
CMP1397 simr-1(cmp291[simr-1::3xHA::TurboID]) I
CMP1411
simr-1(cmp291[simr-1::3xHA::TurboID]) I; bpl-1(cmp292[bpl-
1::degron] ) II
CMP1427
simr-1(cmp291[simr-1::3xHA::TurboID]) I; bpl-1(cmp292[bpl-
1::degron] ) II; ieSi57[Peft-3::TIR1::mRuby::unc-54 3' UTR] II
CMP1428
simr-1(cmp291[simr-1::3xHA::TurboID]) I; bpl-1(cmp292[bpl-
1::degron] ) II; ieSi38[Psun-1::TIR1::mRuby::sun-1 3' UTR] IV

Supplementary file 2. C. elegans strain used in the study.

113
Supplementary File 3. Oligonucleotides sequences used in this study.
Additional Information
CACTTGAACTTCAATACGGCAAGATGAGAATGACT
GGAAACCGTACCGCATGCGGTGCCTATGGTAGC
GGAGCTTCACATGGCTTCAGACCAACAGCCTAT
GCTACCATAGGCACCACGAG
ACTCGGGCCAAGCAGAAAATGCTCAAATTTCAGA
TCTCCACTAAGATTCCGAACCAGCTCACCAAACG
CTTCTTTACGTTGAAAATCGCAGAGGCGAAATCAC
TGGCTGAAGGATTCAATATGTACCCATACGACGT
CCC
ATATACACTAGTGGAATGCGGGAGTAAATTATAAA
AGTTTGAAGGGGAGTATGCATTTATAATTTGGGAG
AAAACTATGGAATCTAGGGTAAAACGAAAGTCATA
CAAGAAATGTCTCTACTTCTCGGCGGAACGAAGG
G
CGCGCAAGTATCCGATGATA
TCAACTCCCCAATCACGAAG
CGCAGAGGCGAAATCACTGG
CATAAATATTAGGATTGTAG
ATGATGAAGGGTCTAATTCGTCATAAATATCCTAA
AGATCCAGCCAAACCTCCGGCCAAGGCACAAGTT
GTGGGATGGCCACCGGTGAGATCATACCGGAAG
AACGTGATGGTTTCCTGCCAAAAATCAAGCGGTG
GCCCGGAGGCGGCGGCGTTCGTGAAGTAGGATT
GTAGCGGATACAGATAGATCAGTGTATAAAT
CCTAAAGAAGCCGAAACACG
TTGAGACGGTTGTGGATGAG
Identifiers
co-CRISPR donor
oligo
guide RNA oligo
PCR amplification
PCR amplification
genotyping primer
genotyping primer
guide RNA oligo
guide RNA oligo
repair template
genotyping primer
genotyping primer
Designation
dpy-10(cn64) donor oligo
dpy-10(cn64) guide RNA
simr-1- TurboID primer - F
simr-1- TurboID primer - R
simr-1 TurboID primer - F
simr-1 TurboID primer - R
simr-1 guide RNA
bpl-1 guide RNA
bpl-1 repair template
bpl-1 genotyping primer - F
bpl-1 genotyping primer - R
Reagent type
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent

Supplementary file 3. Oligonucleotides used in this study.
114
Chapter IV
Identifying RNA modifying proteins at multiple steps in the C. elegans
RNA silencing pathway

Introduction
RNA interference (RNAi) is an evolutionarily conserved silencing mechanism that
is critical in protecting the C. elegans genome from deleterious and/or foreign RNAs
(Phillips et al., 2012; Timmons and Fire, 1998; Vella and Slack, 2005; Wolfswinkel et al.,
2009). The three major classes of C. elegans noncoding small RNAs include small
interfering RNAs (siRNAs), microRNAs (miRNAs) and PIWI-interacting RNAs (piRNAs).
They are typically 18-30 nucleotides long and are categorized based on their length and
5’ nucleotide (Zhang et al., 2011). siRNAs can be grouped into two subclasses,
exogenous and endogenous. Exogenous siRNAs originate from double stranded RNA
(dsRNA) taken up from the environment while endogenous siRNAs are derived from
transposons, coding genes and aberrant transcripts (Phillips et al., 2012; Zhang et al.,
2011). The mechanisms for identification of RNA to be silenced by siRNAs can vary, but
one such mechanism is the recognition of dsRNA by Dicer, a conserved RNase III
enzyme (Figure 1A)(Duchaine et al., 2006; Ketting et al., 2001; Phillips et al., 2012). Upon
recognition, Dicer cleaves the dsRNA, generating primary siRNAs which act as a guide
for the Argonaute protein RDE-1 to silence complementary mRNA transcripts (Carmell et
al., 2002). The mechanisms by which Argonaute proteins mediate silencing can vary,
from direct cleaving, decapping and deadenylating the target transcripts or translation
inhibition (Gu et al., 2009; Kennedy et al., 2004).
115
Primary siRNAs are generally present at low levels , necessitating production of
secondary siRNAs by the mutator complex (Pak and Fire, 2007; Phillips et al., 2012; Yigit
et al., 2006). Within the mutator complex, the RNA-dependent RNA polymerase (RdRP)
RRF-1 synthesizes secondary siRNAs that are typically 22nt in length and contain a 5’-
triphosphorylated guanosine (22G-RNAs)(Gu et al., 2012; Sijen et al., 2007; Tsai et al.,
2015). These newly amplified secondary siRNAs can be bound by worm-specific
Argonaute (WAGO) proteins downstream of the amplification step to promote silencing
of target mRNAs (Figure 1A). This amplification by RdRP amplifies the strength of the
silencing signal and also facilitates inheritance of silencing in subsequent generations.
The mutator complex forms perinuclear foci that are germline specific, and contains a
collection of proteins critical to secondary siRNA amplification, whose localization and
interactions are mediated by the protein MUT-16. mut-16 knockout leads to a loss of
Mutator foci and thus mislocalization of all mutator proteins as well as a severe decrease
in secondary siRNA production and in turn, a loss of robust silencing (Phillips et al., 2012).
The mechanism of biogenesis of these secondary siRNAs and what factors
determine their exact length of 22 nucleotides is not fully understood. It is possible that
the processivity of RRF-1 is sufficient to generate siRNAs of precisely 22 nucleotides, but
we favor the hypothesis that RRF-1 generates longer siRNA precursors and a 3’-5’
exonuclease is then required to the generate siRNAs of the correct size (Figure 1B). This
model is supported by evidence of 3’ small RNA trimming in a variety of RNA silencing
pathways, in numerous organisms. Members of the PARN 3’-5’ exonuclease family,
PARN-1 and PNLDC1 in C. elegans and B. mori respectively, have been found to trim
the 3’ ends of piRNAs while Nibbler, a 3’-5’ exoribonuclease in D. melanogaster, is
116
involved in miRNA 3’ end trimming (Han et al., 2011; Izumi et al., 2016; Tang et al., 2016).
These findings suggest that despite differences between small RNA types and between
species, the mechanisms for small RNA trimming may be conserved. Without proper
processing, the amplification of the silencing signal will be lost as WAGO proteins have
specific binding requirements with regard to siRNA length. Understanding this aspect of
RNA silencing can have implications in studies of cancer, infertility and other diseases
related to genome stability.
In tandem with understanding secondary siRNA 3’ end processing, characterizing
the mechanism by which mRNA targeted by RNA silencing are modified for use as
templates in secondary siRNA amplification is necessary. The mutator complex,
aforementioned site of siRNA amplification, contains many proteins that have been
identified but remain incompletely characterized (Phillips et al., 2012). MUT-16, the core
of the mutator complex, recruits other proteins involved in amplification, including the 3’-
5’ exonuclease MUT-7, the DEAD-box RNA helicases MUT-14 and SMUT-1, and two
uncharacterized proteins, RDE-2 and MUT-15 as well as others (Phillips et al., 2012;
Uebel et al., 2018). One of these proteins that has been shown to be required for
production of mutator-dependent siRNAs is MUT-2, a nucleotidyltransferase (Chen et al.,
2005; Gu et al., 2009; Phillips et al., 2012; Zhang et al., 2011).
MUT-2, based on its amino acid sequence, is predicted to be a homolog of the b
nucleotidyltransferase family, many of which modify endo-siRNAs, miRNAs, or miRNA
precursors (Chen et al., 2005; Heo et al., 2009; Wolfswinkel et al., 2009). Proteins in this
superfamily contain a nucleotidyltransferase domain that consists of a conserved glycine-
serine motif and aspartic-acid triad. This aspartic-acid triad coordinates two divalent metal
117
cations involved in a two-metal ion mechanism of nucleotide addition (Martin et al., 2004).
This nucleotidyltransferase domain is commonly found in poly (A) polymerases and
terminal uridyl transferases. Mutations compromising these two features leads to an RNAi
deficient phenotype (Chen et al., 2005). Those same residues are conserved in the fission
yeast homolog, Cid12, which associates with RdRPs and is required for S. pombe siRNA
accumulation (Chen et al., 2005).
So far, analysis of homologous proteins or proteins that share similar functional
domains have been suggestive of MUT-2 involvement in uridylation (Figure 1C)(Han et
al., 2011; Kwak and Wickens, 2007; Liu et al., 2011; Marasovic et al., 2013; Wolfswinkel
et al., 2009). However, small RNA sequencing data in C. elegans has shown that it is
likely not involved in tailing of siRNAs (Phillips et al., 2014). This suggests MUT-2 function
is critical to the production of siRNAs but is not directly related to the processing of those
small RNAs.
Here, we perform an RNAi screen to identify candidate exonucleases involved in
trimming of 22G-RNAs during secondary siRNA amplification at the mutator complex.
Additionally, we began examination of MUT-2 and the role it plays in the modification of
mRNA prior to use as a template for siRNA amplification. This work is important as the
details of small RNA and mRNA modification in C. elegans silencing pathways, is not well
understood.

118
Results
3’ Exonuclease candidates did not change siRNA length
In order to address the question of whether a previously unidentified protein is
involved in 3’ end trimming of nascent 22G siRNAs, a list of candidate genes containing
3’-5’ exonuclease domains was generated through Wormbase, utilizing both the Pfam
and Interpro protein databases (Blum et al., 2020; Mistry et al., 2020). This search
returned a list of candidate exonucleases (Supplementary file 1), with which I then
conducted an RNAi screen of wild-type animals fed RNAi clones targeting each
candidate. After two generations of exposure to RNAi, to ensure full silencing of the
candidate proteins, gravid animals were collected and total RNA was isolated using Trizol,
followed by chloroform extraction and isopropanol precipitation. For candidate genes
where two generations of RNAi resulted in lethality or sterility, worms were raised for only
a single generation on RNAi prior to RNA isolation (Supplementary file 1). Additionally,
for exonuclease candidates where mutant animals were readily available, in parallel I
isolated RNA from wild-type and mutant adult animals (Supplementary file 1).
Following RNA isolation, I performed Northern blots for a WAGO-class 22G-RNA
by running the samples on a polyacrylamide gel, transferring to a nylon membrane and
probing with a digoxigenin (DIG)-labeled locked nucleic acid (LNA) probe (See Materials
and Methods and Table 1). This non-radioactive Northern protocol utilizes an alkaline
phosphatase-conjugated DIG antibody as a safer alternative to radiolabeled LNA probes.
I predicted that RNAi knockdown or mutants of exonucleases that are WAGO 22G-RNA
trimming, would display an increase in siRNA length (Figure 2), however no substantial
changes in siRNA length were detected. I did observe that mutants in mut-7, rde-10, and
119
eri-1 resulted in loss of the WAGO-class 22G-RNA, which is expected as these three
genes were previously known to play a role in RNA silencing pathways (Phillips
2012,Uebel 2018).
To determine whether any of these exonucleases are involved in miRNA or piRNA
3’ trimming, I similarly performed Northern blotting with RNA from the RNAi knockdown
or exonuclease mutants, and probed for individual miRNAs and piRNAs. I found no
changes to miRNA length, but single deletions in parn-1 and parn-2 resulted in increased
length of piRNAs, though curiously, the RNAi knockdowns and the parn-1 parn-2 had no
effect (Figure 2- figure supplement 1). PARN family proteins have been shown to be
involved in 3’ end piRNA trimming, demonstrating that the non-radioactive Northern
protocol was capable of capturing changes in small RNA length (Tang 2016). Though the
identification of changes to small RNA length is feasible through the use of the non-
radioactive Northern protocol, it appears that RNAi of candidate exonucleases may not
have been sufficient to cause a change in small RNA length.

Catalytic activity is required for MUT-2 function but not localization
Next, to examine modifications made to mRNA during RNA silencing, I chose to
study the nucleotidyltransferase MUT-2. I generated a catalytically dead mut-2 animal
(mut-2(cd)::gfp) by mutating two aspartic acid residues (D105 and D107) into alanine in
key positions of the MUT-2 nucleotidyltransferase catalytic domain using CRISPR/Cas9
technology (Figure 3A). To assess whether catalytic activity is necessary for effective
exogenous RNAi, I exposed mut-2(cd)::gfp , mut-2::gfp, mut-16(-), and wild-type animals
to somatic (lin-29 or nhr-23) and germline (pos-1) RNAi. mut-2::gfp elicited an RNAi
120
phenotype similar to wild-type animals, while mut-2(cd)::gfp animals displayed a RNAi
defective phenotype, similar to the RNAi defect observed previously in mut-16(-) animals
(Figure 3B) (Phillips et al., 2012, Figure 1B). These data indicate that MUT-2
nucleotidyltransferase activity is necessary for its function as a critical part of the RNA
silencing pathway.
To determine if the catalytic domain of MUT-2 is necessary for its localization to
the mutator complex, I performed immunofluorescence on mut-2::gfp and mut-2(cd)::gfp
strains. Both wild-type and catalytically dead MUT-2 localized normally to Mutator foci
(Figure 3C), indicating that catalytic activity is not required for MUT-2 localization. Taken
together, this suggests that the catalytic activity of MUT-2 is essential to its function as
part of the RNA silencing pathway, but is not critical to its association with the mutator
complex or localization to Mutator foci.

3’ End-Seq protocol development for MUT-2 characterization
To determine whether MUT-2 modifies the 3’ ends of small RNA-targeted mRNAs,
I worked to optimize and troubleshoot a 3’ End Sequencing (3’ End-Seq) method, which
would allow for sequencing of mRNA 3’ ends, including non-templated modifications. I
utilized an siRNA sensor strain, ubl-1::GFP-siR-1-sensor where a target site for an
abundant, endogenous siRNA, 22G siR-1, triggers silencing of the GFP transcript
(Montgomery et al., 2012). However, if the RNAi pathway is disrupted, the GFP transcript
is no longer silenced and becomes visible throughout the animal. The sensor strain, in
the presence or absence of the mut-2 mutant allele, were plated on lethal pos-1 RNAi
(Figure 4A). pos-1 RNAi and the ubl-1::GFP-siR-1-sensor strain were used for protocol
121
development as they each have clear phenotypes indicative of effective RNAi knockdown.
Based on the presence of GFP expression and progeny viability on pos-1 RNAi, I was
able to confirm RNAi disruption in the mut-2; ubl-1::GFP-siR-1-sensor strain, while ubl-
1::GFP-siR-1-sensor animals were fully RNAi competent. Use of pos-1 RNAi and ubl-
1::GFP-siR-1-sensor strain also provide two potential targets, pos-1 or GFP mRNA,
during library construction optimization, though initially I focused on pos-1.
After RNA isolation from each strain by Trizol, followed by chloroform extraction
and isopropanol precipitation, I began optimization of library construction (Figure 4B).
Following DNase treatment, I used T4 RNA ligase to ligate on a 3’ adaptor which was
then used for adaptor-specific reverse transcription to generate cDNA. A first round of
polymerase chain reaction (PCR) was conducted using a pos-1 specific forward primer
and a reverse transcription (RT) primer. During the second round of PCR, a more internal
pos-1 specific primer bound to an Illumina adaptor was utilized as well as indexed RT
primers. The completed library could then be sent out for sequencing analysis (Figure
4B).
In order to test the gene-specific forward primers, primer one was used for PCR
round one in conjunction with the RT primer. Subsequently, all five pos-1 primers were
tested for PCR round two to ensure that the internal gene-specific primers were
functional, which yielded a cascading sequence of PCR products (Figure 4C,
Supplemental file 3). This suggested that any gene-specific forward primer in combination
with the reverse transcription primer could be used for the PCR amplification steps in
library preparation. However, during PCR round two, an abundance of off target PCR
products were present, most noticeably a consistent product present in both wild-type
122
and mut-2 backgrounds at ~750 nucleotides. In order to minimize off-target products,
various annealing temperatures were tested as well as utilizing new RT primers
containing a small anchor sequence of repeating nucleotides (Supplemental file 3) to
ideally increase specificity. However, this seemed to have no effect on non-specific
amplification (data not shown). Additionally, the amount of PCR product per reaction was
significantly different between WT and mut-2 strains, likely caused by mRNA
concentrations being low in the wild-type background as RNAi was functioning normally
(Figure 4C). Additional PCR cycles may be necessary during library construction.

Discussion
RNAi is a critical silencing mechanism, and identifying the protein(s) involved in
siRNA 3’ end trimming as well as how mRNA is modified before secondary siRNA
biogenesis occurs has widespread implications. As biogenesis of 22G tri-phosphorylated
siRNAs is a critical step in the amplification of this silencing signal, fully understanding
the mechanism by which this transcript production is occurring is of utmost importance.
Loss of this ability to properly propagate the silencing signal can lead to increased
susceptibility to many forms of DNA damage, and understanding this silencing
mechanism may shed light on some illnesses seen in humans involving genome
instability caused by a weakened silencing response.

3’ Exonuclease mutants may be necessary to observe changes in siRNA length
Though there were no clear indicators of a loss of 3’ end trimming during RNAi of
the exonuclease candidates, there are several possibilities for further analysis. The 22G-
123
siR1 LNA probe used during Northern blot hybridization seemed to identify several bands
of various sizes for each sample. Due to the qualitative and preliminary nature of this
analysis, determination of changes in relative levels of each band and whether loss of the
RNAi targeted nuclease led to shifts from one band length to another was not possible.
The lack of a small RNA ladder also did not allow for identification of the actual length of
siRNAs populating each sample. Through a combination of a more quantitative approach
and more accurate siRNA length determination, a second look at each exonuclease
candidate is warranted. Ideally, future work down this line of inquiry will utilize mutant
animals instead of RNAi. Based on the genetic mutants of eri-1 and rde-10 mutants
showing a loss of the 22G-siR1, but the RNAi of the same genes having no effect, RNAi
knockdown may not be the most efficient or effective way to disrupt siRNA processing.
Utilizing mutants in future experiments will likely lead to a more accurate representation
of the small RNA profile, as evidenced by the piRNA length shift in PARN protein family
mutants (Figure 2- figure supplement 1). However, the lack of change in piRNA length for
the parn-1/2 double mutant is concerning, potentially indicating an issue with the mutant
itself.
Additionally, this preliminary work ruled out many known proteins containing 3’-5’
exonuclease domains as candidates for siRNA 3’ end trimming but it is possible that any
protein containing an exonuclease domain can potentially be involved in this process. It
is also possible that exonuclease activity is not needed for small RNA trimming and that
some other mechanism is utilized to ensure proper 22-base length in newly synthesized
siRNAs.

124

MUT-2 is involved in making poly(UG) additions to mRNA
Following optimization of 3’-End Seq for use in identifying changes in mRNA
modifications during pos-1 RNAi in mut-2 mutant animals, the next step was to generate
libraries and send them out for sequencing. However, another research group has
subsequently shown that MUT-2 is involved in adding stretches of non-templated uridine
and guanosine ribonucleotides to the 3’ ends of RNA targeted during transposon silencing
and transgenerational epigenetic inheritance (Shukla et al., 2020). These poly(UG) or
pUG tails are added to germline and soma expressed RNAs and it has been suggested
that these pUG tails help to stabilize mRNA fragments and possibly form a structure
essential for RdRP recruitment. During RNAi induced silencing, these pUG RNAs persist
for several generations, but are not permanent. Over repeated cycles of siRNA synthesis,
pUG RNAs shorten during each round, possibly as a result of RdRP-dependent
secondary siRNA synthesis. However, the natural targets of pUG tailing such as
transposons, are silenced indefinitely, suggesting that endogenous targets of pUG tailing
are reinforced at each generation (Shukla et al., 2020). The findings by the Kennedy lab
align with our own initial hypotheses of MUT-2 activity involving the addition of uridine
residues to 3’ ends of mRNA, though their work provides a more interesting reality with
the additions being alternating uridine and guanosine ribonucleotides.

125
Materials and Methods
Strains
The wild-type C. elegans strain used was N2. Worms were raised at 20°C under
standard conditions (Brenner, 1974). All strains used for this project are listed in
Supplementary file 2.

Plasmid and strain construction
unc-119(ed3) III; mgSi4[pCMP2(ubl-1p::GFP::siR-1-sensor-ubl-1-3'UTR Cbr-unc-
119(+))] IV , mut-2(ne298) I; mgSi4 IV and mut-2::gfp::3xFLAG were designed previously
(Montgomery et al., 2012; Phillips et al., 2012; Uebel et al., 2018). For introduction of the
DD to AA mutation in mut-2::gfp::3xFLAG, an oligo repair template and RNA guide were
used (Supplementary file 3). CRISPR injections were performed according to published
protocols (Dickinson et al., 2013, 2015; Dokshin et al., 2018; Paix, 2015; Ward, 2015).
CRISPR injection mix included 50ng/µl repair oligo, 25ng/µl mut-2 guide RNA plasmid,
50ng/µl eft-3p::cas9::tbb-2 3'UTR (Addgene plasmid # 61251), 25ng/µl pJA42, 20ng/µl
co-injection repair oligo.

Antibody staining and imaging
Immunofluorescence was conducted on one-day old adults (~24 hours after L4),
dissected in egg buffer containing 0.1% Tween-20 and fixed in 1% formaldehyde in egg
buffer as described (Phillips et al., 2009). Dissected gonads were immunostained with
mouse anti-FLAG (M2, Sigma F1804). Alexa -Fluor secondary antibodies were
purchased and utilized from ThermoFisher Scientific. Imaging was conducted on
126
DeltaVision Elite microscope (GE Healthcare) using 60x N.A. 1.42 oil-immersion
objective. Image analysis was performed using SoftWoRx package and when data stacks
were collected, deconvolution was conducted and images were presented as max
intensity projections, followed by pseudo-coloring using Adobe Photoshop.

RNAi Assays
Synchronized L1 worms raised at 20°C and were fed E. coli expressing dsRNA
against pos-1, lin-29 and nhr-23 (Kamath, 2003). For pos-1, embryos were scored for
viability four to five days after parental animals were placed on RNAi. lin-29 and nhr-23
animals were scored three days after initial exposure to RNAi for vulval bursting and larval
arrest, respectively.

DIG-labeled non-radioactive RNA northern protocol
Materials and Reagents
1. 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC; Sigma, cat. No. 7750)
2. 1-Methylimidazole (Sigma, cat. No. M50834)
3. 3MM Whatman chromatography paper (Whatman, cat. No. 3030917)
4. Anti-Digoxigenin-AP, Fab fragments (Roche, cat. No. 11 093 274 910)
5. CSPD (Roche, cat. No. 11 655 884 001)
6. DIG Wash and Block Buffer Set (Roche, cat. No. 11 585 762 001)
7. Nylon Membranes, positively charged (Roche, cat. No. 11 209 299 001)
8. TBE buffer, 10× solution (National diagnostics, cat. No. EC-860)
9. ULTRAhyb Ultrasensitive Hybridization Buffer (Ambion, cat. No. AM8670)
127
10. Exiqon LNA probe- C. elegans- 22G_siR-1
11. Exiqon LNA probe- C. elegans- cel-miRE-1-3p
12. Exiqon LNA probe- C. elegans- 21UR-1

Procedure
A. RNA extraction (1–2 h)
1. Collect 1500 adult stage worms (68 hours post L1 synchronization) on large
plates.
2. Wash worms three times with M9 (H2O acceptable as well).
3. Add 10 volumes Trizol (1 ml).
4. Vortex 1 min.
5. Incubate at room temperature (RT) for 5 min.
6. Store in dry ice 10 min.
7. Thaw at room temp.
8. Vortex 1 min.
9. Pellet worm carcasses for 1 min at 1 krcf.
10. Transfer supernatant to new tube.
11. Add ~0.2 volume chloroform (200 µl), relative to the volume of Trizol used.
Shake vigorously for 20 sec. Let sit at RT for 3 min.
12. Centrifuge for 10 min at 12 krcf (10 krcf if using 15 ml tubes) and 4˚C.
13. Transfer aqueous phase to new tube.
14. Repeat chloroform extraction.
128
15. Add ~0.5 volume isopropanol, relative to the volume of Trizol used (500 µl).
Vortex.
16. Incubate at -20 for 2 hours.
17. Centrifuge for 20 min at 21 krcf and 4˚C.
18. Pour off supernatant.
19. Wash pellet with cold 75% EtOH. (1ml)
20. Spin briefly. (~2 minutes)
21. Pull off as much EtOH as possible.
22. Air dry for a couple minutes, spin for 30 seconds and then pull off any remaining
EtOH.
23. Resuspend pellet in RNAse-free water (50 µl).
24. Normalize to 1.0 µg/µl.
25. Freeze at -80˚C if not continuing on.

B. Prepare denaturing PAGE (1 h)
26. Assemble the electrophoresis device and add running buffer (1x TBE).
27. Rinse the wells of any excess acrylamide and urea with 1× TBE using a syringe
and a needle. Ensure that the wells are clean and devoid of pieces of gel or
urea.
28. Pre-run the gel at 200 V for 30-60 min.

129
C. Sample and marker preparation (0.5 h)
29. Add Ambion Gel Loading Buffer II (equal amount to RNA) to RNA samples
(~10–15 μg RNA) and the size markers. Denature (95 ˚C, 5 min) the RNA
samples and chill the denatured samples on ice.

D. Electrophoresis (1.5–2 h)
30. Immediately before loading samples onto the gel, rinse the wells of the warm
gel thoroughly.
31. Load samples with marker in each wells and run the gel at about 150V for 1
hour 45 min or until bromophenol blue (BPB) from the loading solution reaches
bottom of gel. (BPB and cyanol from the loading solution runs around 15 bases
and 60 bases, respectively.)
32. To enable the estimation of the RNA sizes of the final bands, use a
transilluminator to image the gel along with a ruler placed parallel to the gel.
33. Disassemble the gel apparatus and soak the gel in 1×TBE for 10 min with
shaking.

E. Post Staining with EtBr
34. Stain the gel in 1X TBE containing EtBr for ~5 min.
35. Rinse gel briefly in 1X TBE.
36. Image gel (tRNA – 78nt and 5S rRNA -120 nt) for quality of RNA prep and for
visual loading control

130

F. Transfer gel to membrane (0.5–2 h)
37. Cut nylon membrane and 2 sheets of 3MM Whatman papers to the size of the
gel and soak them in 1×TBE for 5 min.
38. Set up transfer - Black side of tray (-), foam, whatman, gel, Hybond H+
membrane (rinse in H2O and soak in TBE), whatman, foam, red side of tray (+)
39. Roll out bubbles between layers
40. Run 50V for 1 hour in the cold room with ice block in buffer. The majority of the
BPB should remain on the membrane after completion of transfer, an indication
that transfer has occurred.
41. Visualize gel on UV box to check for complete transfer.

G. Cross-linking EDC (1.5–2.5 h)
42. Add 245 l of 12.5 M 1-methylimidazole to 9 ml DEPC-treated water. Adjust pH
to 8.0 with 1 M HCl (usually requires 300 l). This can be prepared 1–2 h
before use and kept at room temperature.
43. Immediately before use, add 0.753 g EDC and make the volume up to 24 ml
with DEPC-treated water. This gives a working solution of 0.16 M EDC in 0.13
M 1-methylimidazole at pH 8 and is sufficient to saturate a 3 MM Whatman
paper cut to size. (Use the EDC cross-linking solution immediately.)
44. Place the damp membrane with RNA side face up onto 3MM Whatman paper
saturated in freshly prepared cross-linking EDC reagent (prepared as
described above).
131
45. Wrap membrane and 3MM Whatman paper in plastic wrap and incubate at 50–
60 °C for 15min–2 h. (As a precaution, the RNA side should not be in direct
contact with the saturated 3MM and excess cross-linking reagent should not
be washed over the RNA face of the membrane.)
46. During the cross-linking, put ultrahyb-oligo buffer and hybridization tube in the
65
o
C hyb-oven to warm up.
47. After cross-linking is complete, rinse membrane in excess RNase-free distilled
water to remove any residual cross-linking solution. Note: The membrane can
be now wrapped in Saran and stored at -20˚C for later use.

H. Hybridization (3–12 h+)
48. After completion of cross-linking, roll membrane with RNA side in and insert
into
hybridization bottle.
49. Add 5 ml of DIG Easy hybridization buffer and pre-hybridize at 37˚C for at least
30 min in hybridization oven (5ml is enough for small bottle).
50. Denature the probe at 95˚C for 1 min and add the probe to the hybridization
buffer to yield a final concentration of 0.5 nM.(1.25µl in 5ml)
51. Hybridize at 37˚C overnight with slow rotation.

I. Washing, blocking, and detection (2–6 h+)
52. Discard hybridization buffer from bottle(dispose of as hazardous waste).
53. Wash the membrane twice with Low Stringent Buffer at 37˚C for 15 min.
132
54. Wash the membrane twice with High Stringent Buffer at 37˚C for 5 min.
55. Briefly rinse the membrane with DIG Washing Buffer at 37˚C for 10 min(Dilute
the washing buffer to 1x before use).
56. Incubate the membrane in Blocking Buffer(Dilute the blocking buffer to 1x using
1x maleic acid-450µl MA+4.05ml H2O+500µl Blocking buff.) for 2 hrs at room
temperature.
57. Replace the blocking buffer in hybridization bottle with DIG antibody solution
prepared by mixing DIG antibody solution and blocking buffer at a ratio of
1:15,000.
58. Incubate the membrane at room temperature for 30 min.
59. Wash the membranes in DIG Washing buffer four times for 15 min each.
60. Incubate membrane in development buffer for 5 min. Note: Do not allow the
membrane to dry if it must be reused to detect additional RNAs.
61. Remove blot from bottle with clean forceps and place on cut sheet protector.
Apply the substrate CPD-Star ready to use solution to the surface of the
membrane, close the sheet protector, incubate for 5 min in the dark(cover blot
evenly).
62. Use roller to lightly remove the excess before transfer of the membrane face
down to a new sheet of saran wrap, seal.
63. Expose the membrane to X-ray films. Note: While a few seconds of exposure
is sufficient to get a strong signal for highly expressed small RNAs, poorly
expressed small RNAs will require more exposure time (siRNAs-10min, 5min,
1min: miRNA-10min, 3-5min
133
J. Stripping blots
64. Bring 250 ml of 0.1% SDS to a boil in the microwave.
65. Pour over used blot, RNA side up, in a glass dish. Shake gently for 20 minutes,
and repeat.

3’ End-Seq Protocol
Materials and Reagents
1. TRizol Reagent (ThermoFisher, cat. No. 15596018)
2. TURBO DNase kit (ThermoFisher, cat. No. AM2238)
3. T4 RNA ligase II truncated (VWR, cat. No. 102715-908)
4. SuperScript III reverse transcriptase (ThermoFisher, cat. No. 18080-044)
5. RNaseOut (ThermoFisher, cat. No. 10777019)

A. TURBO DNase Treatment (Will likely be starting with 10µg RNA at 1.0µg/µl)
1. Add 0.1 volume of 10x TURBO DNase buffer and 1µl of TURBO DNase to the
RNA and mix gently.
2. Incubate at 37°C for 20-30 minutes.
3. Add 0.1 volume of resuspended DNase Inactivation reagent and mix well.
4. Incubate 5 min at room temp, mixing occasionally.
5. Centrifuge at 10,000 x g for 1.5 min, transfer RNA to a fresh tube.
6. Normalize to 1.0µg/µl.

134
B. 3’ ATP-independent ligation
7. Mix the following in PCR tubes (20 ul reactions):
• H20 1 µl
• Total RNA (1µg) 10 µl
• 10X Ligation Buffer 2 µl
• PEG 8000 4 µl
• 100 µM 3’ Adaptor (TruSeq from TAM) 1 µl
• T4 RNA Ligase II truncated (NEB M0373) 1 µl
• RNase OUT 1 µl
8. Incubate at 16°C 16 hours.

C. Trizol:Chloroform extraction
9. Add 10 volumes Trizol (200 µl).
10. Vortex 1 min.
11. Incubate at room temperature (RT) for 5 min.
12. Add ~0.2 volume chloroform (40 µl), relative to the volume of Trizol used.
Shake vigorously for 20 sec. Let sit at RT for 3 min.
13. Centrifuge for 10 min at 12 krcf at 4˚C.
14. Transfer aqueous phase to new tube.
15. Repeat chloroform extraction.
16. Add ~0.5 volume isopropanol, relative to the volume of Trizol used (100 µl) and
glycogen (1µl). Vortex.
17. Incubate at RT for 20 mins.
135
18. Centrifuge for 20 min at 21 krcf and 4˚C.
19. Pour off supernatant.
20. Wash pellet with cold 75% EtOH.
21. Spin briefly.
22. Pull off as much EtOH as possible.
23. Air dry for a couple minutes and pull off any remaining EtOH.
24. Resuspend pellet in RNAse-free water (20 µl).

D. Reverse Transcription
25. Combine the following in PCR tubes (set up 1 X 20 µl reactions for each
sample):
• Adaptor-ligated RNA 10 µl
• 100 µM smRNA-seq RT Primer 1 µl
• 10 mM dNTPs 1 µl
26. Heat to 65°C for 5 min. Cool on ice. Spin briefly.
27. Add the following to each reaction:
• 5X First Strand Buffer 4 µl
• 100 mM DTT 2 µl
• RNaseOUT 1 µl
• SuperScriptIII 1 µl
28. Incubate reactions at 50°C for 60 minutes and inactivate reactions at 70°C for
15 minutes. Add 1 µl RNase H (2U) to each reaction and incubate at 37°C for
20 minutes.
136

E. PCR Round 1 (with gene specific forward primer)
29. Combine the following in PCR tubes:
• H2O 34.5 µl
• RT reaction 2.5 µul
• 5X Phusion GC Buffer 10 µl
• 10 µM gene specific F primer 1.0 µl
• 10 µM gene TruSeq R primer 1.0 µl
• 25 mM dNTP mix 0.5 µl
• Phusion polymerase 0.5 µl
30. PCR:
• 98°C for 1 min
• 98°C for 10 sec à60°C for 30 sec à 72°C for 15 sec; ~15 cycles
• 72°C for 5 min
31. Size select 300-600bp amplicons on 8% polyacrylamide gel

F. PCR Round 2 (addition of Illumina adaptors)
32. Combine the following in PCR tubes:
• H2O 34.5 µl
• PCR reaction 1 2.5 µl
• 5X Phusion GC Buffer 10 µl
• 10 µM internal gene specific F 1.0 µl
• 10 µM gene TruSeq R primer 1.0 µl
137
• 25 mM dNTP mix 0.5 µl
• Phusion polymerase 0.5 µl
33. PCR:
• 98°C for 1 min
• 98°C for 10 sec à60°C for 30 sec à 72°C for 15 sec; 14 cycles
• 72°C for 5 min
34. Remove primer dimers with AMPure XP beads or gel purification

References
Blum, M., Chang, H.-Y., Chuguransky, S., Grego, T., Kandasaamy, S., Mitchell, A., Nuka,
G., Paysan-Lafosse, T., Qureshi, M., Raj, S., et al. (2020). The InterPro protein families
and domains database: 20 years on. Nucleic Acids Res 49, gkaa977-.
Brenner, S. (1974). THE GENETICS OF CAENORHABDITIS ELEGANS. Genetics 77,
71–94.
Carmell, M.A., Xuan, Z., Zhang, M.Q., and Hannon, G.J. (2002). The Argonaute family:
tentacles that reach into RNAi, developmental control, stem cell maintenance, and
tumorigenesis. Genes & Development 16, 2733–2742.
Chen, C.-C.G., Simard, M.J., Tabara, H., Brownell, D.R., McCollough, J.A., and Mello,
C.C. (2005). A Member of the Polymerase β Nucleotidyltransferase Superfamily Is
Required for RNA Interference in C. elegans. Current Biology 15, 378–383.
Dickinson, D.J., Ward, J.D., Reiner, D.J., and Goldstein, B. (2013). Engineering the
Caenorhabditis elegans genome using Cas9-triggered homologous recombination. Nat
Methods 10, 1028–1034.
Dickinson, D.J., Pani, A.M., Heppert, J.K., Higgins, C.D., and Goldstein, B. (2015).
Streamlined Genome Engineering with a Self-Excising Drug Selection Cassette. Genetics
200, 1035–1049.
Dokshin, G.A., Ghanta, K.S., Piscopo, K.M., and Mello, C.C. (2018). Robust genome
editing with short single-stranded and long, partially single-stranded DNA donors in C.
elegans. Biorxiv 352260.
138

Duchaine, T.F., Wohlschlegel, J.A., Kennedy, S., Bei, Y., Jr, D.C., Pang, K., Brownell,
D.R., Harding, S., Mitani, S., Ruvkun, G., et al. (2006). Functional Proteomics Reveals
the Biochemical Niche of C. elegans DCR-1 in Multiple Small-RNA-Mediated Pathways.
Cell 124, 343–354.
Gu, W., Shirayama, M., Conte, D., Vasale, J., Batista, P.J., Claycomb, J.M., Moresco,
J.J., Youngman, E.M., Keys, J., Stoltz, M.J., et al. (2009). Distinct argonaute-mediated
22G-RNA pathways direct genome surveillance in the C. elegans germline. Molecular
Cell 36, 231–244.
Gu, W., Lee, H.-C., Chaves, D., Youngman, E.M., Pazour, G.J., Jr, D.C., and Mello, C.C.
(2012). CapSeq and CIP-TAP Identify Pol II Start Sites and Reveal Capped Small RNAs
as C. elegans piRNA Precursors. 151, 1488–1500.
Han, B.W., Hung, J.-H., Weng, Z., Zamore, P.D., and Ameres, S.L. (2011). The 3’-to-5’
exoribonuclease Nibbler shapes the 3’ ends of microRNAs bound to Drosophila
Argonaute1. Current Biology : CB 21, 1878–1887.
Heo, I., Joo, C., Kim, Y.-K., Ha, M., Yoon, M.-J., Cho, J., Yeom, K.-H., Han, J., and Kim,
V.N. (2009). TUT4 in Concert with Lin28 Suppresses MicroRNA Biogenesis through Pre-
MicroRNA Uridylation. Cell 138, 696–708.
Izumi, N., Shoji, K., Sakaguchi, Y., Honda, S., Kirino, Y., Suzuki, T., Katsuma, S., and
Tomari, Y. (2016). Identification and Functional Analysis of the Pre-piRNA 3′ Trimmer in
Silkworms. Cell 164, 962–973.
Kamath, R. (2003). Genome-wide RNAi screening in Caenorhabditis elegans. Methods
30, 313–321.
Kennedy, S., Wang, D., and Ruvkun, G. (2004). A conserved siRNA-degrading RNase
negatively regulates RNA interference in C. elegans. Nature 427, 645–649.
Ketting, R.F., Fischer, S.E., Bernstein, E., Sijen, T., Hannon, G.J., and Plasterk, R.H.
(2001). Dicer functions in RNA interference and in synthesis of small RNA involved in
developmental timing in C. elegans. Genes & Development 15, 2654–2659.
Kwak, J.E., and Wickens, M. (2007). A family of poly(U) polymerases. RNA (New York,
N.Y.) 13, 860–867.
Liu, N., Abe, M., Sabin, L.R., Hendriks, G.-J., Naqvi, A.S., Yu, Z., Cherry, S., and Bonini,
N.M. (2011). The Exoribonuclease Nibbler Controls 3′ End Processing of MicroRNAs in
Drosophila. Current Biology 21, 1888–1893.
139
Marasovic, M., Zocco, M., and Halic, M. (2013). Argonaute and Triman Generate Dicer-
Independent priRNAs and Mature siRNAs to Initiate Heterochromatin Formation.
Molecular Cell 52, 173–183.
Martin, G., Möglich, A., Keller, W., and Doublié, S. (2004). Biochemical and Structural
Insights into Substrate Binding and Catalytic Mechanism of Mammalian Poly(A)
Polymerase. Journal of Molecular Biology 341, 911–925.
Mistry, J., Chuguransky, S., Williams, L., Qureshi, M., Salazar, G.A., Sonnhammer,
E.L.L., Tosatto, S.C.E., Paladin, L., Raj, S., Richardson, L.J., et al. (2020). Pfam: The
protein families database in 2021. Nucleic Acids Res 49, gkaa913-.
Montgomery, T.A., Rim, Y.-S., Zhang, C., Dowen, R.H., Phillips, C.M., Fischer, S.E.J.,
and Ruvkun, G. (2012). PIWI Associated siRNAs and piRNAs Specifically Require the
Caenorhabditis elegans HEN1 Ortholog henn-1. 8, e1002616.
Paix, A. (2015). High Efficiency, Homology-Directed Genome Editing in Caenorhabditis
elegans Using CRISPR-Cas9 Ribonucleoprotein Complexes. Genetics Investigation 1–
26.
Pak, J., and Fire, A. (2007). Distinct populations of primary and secondary effectors
during RNAi in C. elegans. Science 315, 241–244.
Phillips, C.M., McDonald, K.L., and Dernburg, A.F. (2009). Cytological Analysis of Meiosis
in Caenorhabditis elegans. pp. 171–195.
Phillips, C.M., Montgomery, T.A., Breen, P.C., and Ruvkun, G. (2012). MUT-16 promotes
formation of perinuclear mutator foci required for RNA silencing in the C. elegans
germline. Genes & Development 26, 1433–1444.
Phillips, C.M., Montgomery, B.E., Breen, P.C., Roovers, E.F., Rim, Y.-S., Ohsumi, T.K.,
Newman, M.A., Wolfswinkel, J.C. van, Ketting, R.F., Ruvkun, G., et al. (2014). MUT-14
and SMUT-1 DEAD Box RNA Helicases Have Overlapping Roles in Germline RNAi and
Endogenous siRNA Formation. Current Biology 24, 839–844.
Shukla, A., Yan, J., Pagano, D.J., Dodson, A.E., Fei, Y., Gorham, J., Seidman, J.G.,
Wickens, M., and Kennedy, S. (2020). poly(UG)-tailed RNAs in genome protection and
epigenetic inheritance. Nature 1–24.
Sijen, T., Steiner, F.A., Thijssen, K.L., and Plasterk, R.H.A. (2007). Secondary siRNAs
result from unprimed RNA synthesis and form a distinct class. Science 315, 244–247.
Tang, W., Tu, S., Lee, H.-C., Weng, Z., and Mello, C.C. (2016). The RNase PARN-1 Trims
piRNA 3’ Ends to Promote Transcriptome Surveillance in C. elegans. Cell 164, 974–984.
140
Timmons, L., and Fire, A. (1998). Specific interference by ingested dsRNA. Nature 395,
854–854.
Tsai, H.-Y., Chen, C.-C.G., Jr, D.C., Moresco, J.J., Chaves, D.A., Mitani, S., III, J.R.Y.,
Tsai, M.-D., and Mello, C.C. (2015). A Ribonuclease Coordinates siRNA Amplification
and mRNA Cleavage during RNAi. Cell 160, 407–419.
Uebel, C.J., Anderson, D.C., Mandarino, L.M., Manage, K.I., Aynaszyan, S., and Phillips,
C.M. (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 Genetics 14, e1007542-22.
Vella, M.C., and Slack, F.J. (2005). C. elegans microRNAs. WormBook : The Online
Review of C. Elegans Biology 1–9.
Ward, J.D. (2015). Rapid and Precise Engineering of the Caenorhabditis elegans
Genome with Lethal Mutation Co-Conversion and Inactivation of NHEJ Repair. Genetics
199, 363–377.
Wolfswinkel, J.C. van, Claycomb, J.M., Batista, P.J., Mello, C.C., Berezikov, E., and
Ketting, R.F. (2009). CDE-1 Affects Chromosome Segregation through Uridylation of
CSR-1-Bound siRNAs. Cell 139, 135–148.
Yigit, E., Batista, P.J., Bei, Y., Pang, K.M., Chen, C.-C.G., Tolia, N.H., Joshua-Tor, L.,
Mitani, S., Simard, M.J., and Mello, C.C. (2006). Analysis of the C. elegans Argonaute
Family Reveals that Distinct Argonautes Act Sequentially during RNAi. Cell 127, 747–
757.
Zhang, C., Montgomery, T.A., Gabel, H.W., Fischer, S.E.J., Phillips, C.M., Fahlgren, N.,
Sullivan, C.M., Carrington, J.C., and Ruvkun, G. (2011). mut-16 and other mutator class
genes modulate 22G and 26G siRNA pathways in Caenorhabditis elegans. Proceedings
of the National Academy of Sciences of the United States of America 108, 1201–1208.

141
Figures and Figure Legends

Figure 1. RNA silencing is strengthened at the mutator complex.
(A) Biogenesis of 22G secondary siRNAs occurs when primary siRNAs generated from exogenous sources are
amplified by a collection of proteins localized to the mutator complex to produce secondary siRNAs. These secondary
siRNA can then be utilized by secondary worm-specific Argonaute (WAGO) proteins to robustly silence mRNA targets.
(B) The presence or absence of a 3’-5’ exonuclease during secondary siRNA production is currently unknown. (C)
MUT-2 is a known component of the mutator complex but its specific role was unclear.

AAAAA
Primary
AGO
Dicer
Primary
siRNAs
RdRP
mutator
complex
Secondary
siRNAs
Primary
AGO
Secondary
AGO Secondary
AGO
Secondary
AGO
Secondary
AGO
Secondary
AGO
Secondary
AGO
A
RdRP
3’-5’
Exonuclease?
mRNA
template
B
RdRP
mRNA
template
C
MUT-2?
142

Figure 2. 3’-5’ exonuclease knockdown showed no significant change to siRNA length.
(A) Northern blot exposures of 3’-5’ exonuclease candidates identified using LNA detection probe targeting 22G siR-1
small RNAs.

A
ccr-4
C01B10.3
Y56A3A.33
exos-1
ZK1098.3
exos-9
panl-2
disl-2
B0393.4
xrn-2
parn-2
Y57A10A.13
crn-5
crn-3
F07H5.10
ZK1248.15
20nt -
30nt -
crn-4
exos-7
dcs-1
parn-1
C08B6.8
hint-3
C05C8.5
gon-14
F14F9.5
K10C9.3
exos-4.1
dis-3
egal-1
M02B7.2
R02D3.8
eri-1
20nt -
30nt -
siRNA siRNA
F30A10.0
W05H12.2
W04A8.4
rde-10
xrn-1
dcap-1
dcap-2
dcap-1/2
wild-type (no RNAi)
rde-10(hj20)
parn-1(tm869)
parn-1(Rb1621)
parn-2(tm1339)
mut-7(pk204)
ZK1098.3
eri-1(mg366)
20nt -
30nt -
siRNA
dom-3
C37H5.14
ccf-1
mut-7
angl-1
L4440
Y113G7B.12
exos-2
BE0003N10.1
smg-6
exos-8
Y17G7B.12
mut-7(pk720)
wild-type (no RNAi)
pqe-1
eol-1
20nt -
30nt -
siRNA
parn-1/2(tm869/tm1339)
143

Figure 2-figure supplement 1. miRNA and piRNA probes targeting 3’-5’ exonuclease candidate small
RNAs.
(A) Northern blot exposures of 3’-5’ exonuclease candidates identified using LNA detection probes targeting miR-
1(miRNA) and 21UR-1(piRNA).
miRNA
crn-4
exos-7
dcs-1
parn-1
C08B6.8
hint-3
C05C8.5
gon-14
F14F9.5
K10C9.3
exos-4.1
dis-3
egal-1
M02B7.2
R02D3.8
eri-1
ccr-4
C01B10.3
Y56A3A.33
exos-1
ZK1098.3
exos-9
panl-2
disl-2
B0393.4
xrn-2
parn-2
Y57A10A.13
crn-5
crn-3
F07H5.10
ZK1248.15
20nt -
30nt -
20nt -
30nt -
20nt -
30nt -
20nt -
30nt -
A
piRNA miRNA piRNA
F30A10.0
W05H12.2
W04A8.4
rde-10
xrn-1
dcap-1
dcap-2
dcap-1/2
wild-type (no RNAi)
rde-10(hj20)
parn-1(tm869)
parn-1(Rb1621)
parn-2(tm1339)
mut-7(pk204)
ZK1098.3
eri-1(mg366)
20nt -
30nt -
20nt -
30nt -
miRNA piRNA
dom-3
C37H5.14
ccf-1
mut-7
angl-1
L4440
Y113G7B.12
exos-2
BE0003N10.1
smg-6
exos-8
Y17G7B.12
pqe-1
mut-7(pk720)
eol-1
wild-type (no RNAi)
20nt -
30nt -
20nt -
30nt -
miRNA piRNA
parn-1/2(tm869
/tm1339)
144

Figure 3. Catalytically dead MUT-2 is RNAi defective but shows no change in localization.
(A) Graphical representation of CRISPR mutagenesis of critical aspartic acid residues in catalytic domain of MUT-2.
(B) Mutation of key residues in MUT-2 catalytic domain leads to RNAi defective phenotype when exposed to germline
and somatic RNAi. (C) Immunofluorescence imaging shows loss of catalytic activity does not change MUT-2 perinuclear
localization. Scale bars, 5µm.

145

Figure 4. 3’ End-Seq protocol development.
(A) Workflow for two generation RNAi exposure prior to 3’ End-Seq library construction. (B) Library construction schema
utilizing gene specific primers with a nested PCR approach. (C) Sequential gene-specific primers are equally effective
during library construction but have some off target amplification.

146

Table 1. Exiqon miRCURY LNA detection probes

C. elegans - 22G_siR-1
Custom Exiqon miRCURY LNA Detection Probe
C. elegans - cel-miR-1-3p
C. elegans - 21UR-1
Sequence (5’-3’)
/5DigN/ACCTCATACCGCGTATCCATT
/5DigN/TACATACTTCTTTACATTCCA
/5DigN/CACGGTTAACGTACGTACCA
147
Supplementary File 1. Proteins targeted by RNAi during 3'-5' exonuclease search
Gene Name Sequence Protein Family
Identifier
Generations
of RNAi
Testing
Method
L4440 - - 2 RNAi
F30A10.9 F30A10.9 INTERPRO:IPR029060 1 RNAi
W05H12.2 W05H12.2 INTERPRO:IPR002562 2 RNAi
W04A8.4 W04A8.4 INTERPRO:IPR002562 2 RNAi
F07H5.10 F07H5.10 PFAM:PF08652 2 RNAi
CRN-3 C14A4.4 INTERPRO:IPR002562 1 RNAi
CRN-5 C14A4.5 PFAM:PF01138 1 RNAi
Y57A10A.13 Y57A10A.13 INTERPRO:IPR002562 2 RNAi
XRN-2 Y48B6A.3 INTERPRO:IPR004859 1 RNAi
B0393.4 B0393.4 PFAM:PF13017 2 RNAi
DISL-2 F48E8.6 PFAM:PF00773 2 RNAi
PANL-2 F31E3.4 INTERPRO:IPR012337 2 RNAi
EXOS-9 F37C12.13 PFAM:PF01138 2 RNAi
EXOS-1 Y48A6B.5 PFAM:PF10447 2 RNAi
Y56A3A.33 Y56A3A.33 INTERPRO:IPR012337 2 RNAi
R02D3.8 R02D3.8 INTERPRO:IPR012337 2 RNAi
M02B7.2 M02B7.2 INTERPRO:IPR012337 2 RNAi
EGAL-1 C10G6.1 INTERPRO:IPR002562 2 RNAi
DIS-3 C04G2.6 PFAM:PF00773 2 RNAi
EXOS-4.1 B0564.1 PFAM:PF01138 2 RNAi
K10C9.3 K10C9.3 PFAM:PF00445 2 RNAi
F14F9.5 F14F9.5 PFAM:PF03372 2 RNAi
GON-14 F44C4.4 INTERPRO:IPR012337 2 RNAi
C05C8.5 C05C8.5 INTERPRO:IPR012337 2 RNAi
HINT-3 C26F1.7 PFAM:PF11969 2 RNAi
C08B6.8 C08B6.8 INTERPRO:IPR012337 2 RNAi
DCS-1 Y113G7A.9 INTERPRO:IPR011145 2 RNAi
EXOS-7 F31D4.1 PFAM:PF01138 2 RNAi
CRN-4 AH9.2 INTERPRO:IPR012337 2 RNAi
Y17G7B.12 Y17G7B.12 INTERPRO:IPR012337 2 RNAi
EXOS-8 F41G3.14 PFAM:PF01138 2 RNAi
SMG-6 Y54F10AL.2 INTERPRO:IPR029060 2 RNAi
BE0003N10.1 BE0003N10.1 PFAM:PF01138 2 RNAi
EXOS-2 Y73B6BL.3 PFAM:PF01138 1 RNAi
Y113G7B.12 Y113G7B.12 PFAM:PF03372 2 RNAi
ANGL-1 W02G9.5 PFAM:PF03372 2 RNAi
148
C37H5.14 C37H5.14 PFAM:PF08652 2 RNAi
DOM-3 F54C1.2 PFAM:PF08652 2 RNAi
ZK1248.15 ZK1248.15 INTERPRO:IPR029060 2 RNAi
C01B10.3 C01B10.3 PFAM:PF03372 2 RNAi
CCR-4 ZC518.3 PFAM:PF03372 2 RNAi
XRN-1 Y39G8C.1 INTERPRO:IPR004859 2 RNAi
CCF-1 Y56A3A.20 INTERPRO:IPR012337 2 RNAi
DCAP-1 Y55F3AM.12 INTERPRO:IPR036189 2 RNAi
DCAP-2 F52G2.1 INTERPRO:IPR036189 2 RNAi
RDE-10 Y47G6A.4 PFAM:PF13017 2 RNAi/Mutant
PARN-2 Y57A10A.25 INTERPRO:IPR012337 2 RNAi/Mutant
ZK1098.3 ZK1098.3 INTERPRO:IPR002562 2 RNAi/Mutant
ERI-1 T07A9.5 INTERPRO:IPR012337 2 RNAi/Mutant
PARN-1 K10C8.1 INTERPRO:IPR012337 2 RNAi/Mutant
MUT-7 ZK1098.8 INTERPRO:IPR002562 2 RNAi/Mutant
PQE-1 F52C9.8 INTERPRO:IPR012337 2 RNAi/Mutant
EOL-1 T26F2.3 PFAM:PF08652 2 RNAi/Mutant

Supplementary file 1. 3’-5’ exonuclease candidate proteins used in this study.

149
Supplemental file 2. Strains used for this study.
Strain
Designation
Description
N2 wild-type
GR1720 mgSi4[pCMP2(ubl-1p::gfp::siR-1-sensor-ubl-1-3'UTR Cbr-unc-119(+))] IV
GR2009
mut-2(ne298) I; mgSi4[pCMP2(ubl-1p::gfp::siR-1-sensor-ubl-1-3'UTR
Cbr-unc-119(+))] IV
USC852 mut-2(cmp42[(mut-2::gfp + loxP + 3xFLAG)]) I
CMP1168 mut-2(cmp42[(mut-2::GFP + loxP + 3xFLAG)] cmp223[DD to AA])

Supplementary file 2. C. elegans strains used in this study.

150
Supplementary File 3. Oligonucleotides sequences used in this study.
Additional Information
ATCCTCCATATTGTTGACATCTC
ACACGGTTGGAAGCAGTTCCTG
CTCCAAGTTTAACCA
GTGAGACGTCAACAATATGG
AAATCCTTGACTCCCACAGCG
GCCAATCGTGATCCTTGCACAG
TTCAAGACGGCTCTGTGTGATG
GGCATGCTCATACGGTGATCAG
AGATTTGCTCACGGCGTCCATG
GCCTTGGCACCCGAGAATTCCA
AAA
GCCTTGGCACCCGAGAATTCCA
TTT
GCCTTGGCACCCGAGAATTCCA
CCC
GCCTTGGCACCCGAGAATTCCA
GGG
GCCTTGGCACCCGAGAATTCCA
ACAC
TCTTGTCTAGATCACTGTTTTTA
G
AAACCTAAAAACAGTGATCTAG
AC
CGTGGCTATTCAATATCGTGCC
AACTGGAAGTACAGTAACCGGA
TTGGCgACaAAgAAttcaGcgCTAG
cCGTTGCAATTCATATCCCACAA
GCGGCAAGAGTTCTGGAACAA
GAAGAAC
Identifiers
co-CRISPR donor
oligo
guide RNA oligo
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
genotyping primer
guide RNA oligo
guide RNA oligo
repair template
Designation
rol-6(su1006)
rol-6(su1006) – guide RNA
pos-1(1) - 3' End-Seq - F
pos-1(2) - 3' End-Seq - F
pos-1(3) - 3' End-Seq - F
pos-1(4) - 3' End-Seq - F
pos-1(5) - 3' End-Seq - F
TruSeq RT+ A oligo clamp - F
TruSeq RT+ T oligo clamp - F
TruSeq RT+ C oligo clamp - F
TruSeq RT+ G oligo clamp - F
TruSeq RT+ AC oligo clamp - F
DD to AA into mut-2::gfp::3xFLAG -guide
RNA - F
DD to AA into mut-2::gfp::3xFLAG -guide
RNA - R
DD to AA into mut-2::gfp::3xFLAG -
repair template
Reagent type
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent
sequence-based
reagent

Supplementary file 3. Oligonucleotides sequences used in this study.

151
Appendix

SIMR foci are found in the progenitor germ cells of C. elegans
embryos

This manuscript was first published as:
Uebel, C.J., Manage, K.I., and Phillips, C.M. (2021). SIMR foci are found in the
progenitor germ cells of C. elegans embryos. MicroPublication Biology 2021.

Author Contributions
During research into biological liquid-liquid phase separation in the perinuclear
space of germline nuclei, Dr. Celja Uebel noticed the formation of SIMR foci during
embryonic development. She decided to pursue this finding and the resulting research
led to a publication in microPublication Biology. My contributions to this work included
providing experimental reagents as well as contributing to the writing, primarily with
revision and editing.

152

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).

153
Description
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.
154
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
155
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.

Reagents
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]) 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.

156
References

Brenner S. (1974). The Genetics of Caenorhabditis elegans. Genetics 77, 71–94. PMID:
4366476.

Kawasaki I, Shim YH, Kirchner J, Kaminker J, Wood WB, Strome S. (1998). PGL-1, a
predicted RNA-binding component of germ granules, is essential for fertility in C. elegans.
Cell 94(5), 635–45. PMID: 9741628.

Lev I, Rechavi O. (2020). Germ granules allow transmission of small RNA-based parental
responses in the germ plasm. iScience 23, 101831. PMID: 33305186

Manage KI, Rogers AK, Wallis DC, Uebel CJ, Anderson DC, Nguyen DAH, Arca K, Brown
KC, Rodrigues RJC, deAlbuquerque BFM et al. (2020). A Tudor domain protein, SIMR-
1, promotes siRNA production at piRNA-targeted mRNAs in C. elegans. Elife 9, e56731.
PMID: 32338603

Phillips CM, Montgomery TA, Breen PC, Ruvkun G. (2012). MUT-16 promotes formation
of perinuclear Mutator foci required for RNA silencing in the C. elegans germline. Gene
Dev 26, 1433–1444. PMID: 22713602

Seydoux G, Dunn MA. (1997). Transcriptionally repressed germ cells lack a
subpopulation of phosphorylated RNA polymerase II in early embryos of Caenorhabditis
elegans and Drosophila melanogaster. Dev 124, 2191–2201. PMID: 9187145

Strome S, Wood WB. (1982). Immunofluorescence visualization of germ-line-specific
cytoplasmic granules in embryos, larvae, and adults of Caenorhabditis elegans. Proc Natl
Acad Sci 79, 1558–1562. PMID: 7041123

Sundby AE, Molnar RI, Claycomb JM. (2021). Connecting the Dots: Linking
Caenorhabditis elegans small RNA pathways and germ granules. Trends Cell Biol. PMID:
33526340

Uebel CJ, Agbede D, Wallis DC, Phillips CM. (2020). Mutator foci are regulated by
developmental stage, RNA, and the germline cell cycle in Caenorhabditis elegans. G3
Genes Genomes Genetics 10, 3719–3728. PMID: 32763952

Uebel CJ, Phillips CM. (2019). Phase-separated protein dynamics are affected by
fluorescent tag choice. MicroPublication Biology 143. PMID: 32440657

Updike DL, Knutson AK, Egelhofer TA, Campbell AC, Strome S. (2014). Germ-granule
components prevent somatic development in the C. elegans germline. Curr Biol
24(9),970–5. PMID: 24746798

157
Wan G, Fields BD, Spracklin G, Shukla A, Phillips CM, Kennedy S. (2018).
Spatiotemporal regulation of liquid-like condensates in epigenetic inheritance. Nature
557, 679–683. PMID: 29769721

Acknowledgements
We would like to thank members of the Phillips Lab for helpful discussions of the
manuscript.
Funding
This work was supported by the National Institute of Health grants R35 GM119656 (to
C.M.P.), the National Science Foundation Graduate Research Fellowship Program
Grant No. DGE 1418060 (to C.J.U.) and the University of Southern California Research
Enhancement Fellowship (to C.J.U.). C.M.P. 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. The funders had no role in
study design, data collection and analysis, decision to publish, or preparation of the
manuscript.
Author Contributions
C.J.U.: Conceptualization, Investigation, Visualization, Funding Acquisition, Writing -
original draft.
K.I.M.: Resources, Writing – review and editing.
C.M.P.: Supervision, Funding Acquisition, Writing – review and editing.
Reviewed By: Dustin Updike
History: Received February 16, 2021 Accepted February 22, 2021 Published February
22, 2021
Copyright: © 2021 by the authors. This is an open-access article distributed under the
terms of the Creative Commons Attribution 4.0 International (CC BY 4.0) License, which
permits unrestricted use, distribution, and reproduction in any medium, provided the
original author and source are credited.

Abstract (if available)

Abstract The piwi-interacting RNA (piRNA) pathway is an evolutionarily conserved mechanism that plays an important role in the silencing of transposons and other germline genes. Transposons are selfish genetic elements that when unregulated, can damage the integrity of an organism’s genome, impacting future generations and potentially leading to sterility. In Caenorhabditis elegans (C. elegans), regulation of piRNA target genes is mediated by the mutator complex, which generates high levels of secondary siRNAs for these piRNA-target loci to achieve robust silencing. How coordination between piRNA production in P granules and mutator complex-dependent siRNA biogenesis occurs is not well understood. The work presented in this dissertation explores the function of SIMR-1, a Tudor domain protein critical to the coordination between these two important steps in the piRNA pathway. While not necessary for piRNA production, simr-1 mutants fail to produce high levels of secondary siRNAs for many piRNA-target loci suggesting SIMR-1 acts at a step between these two events. It also identifies SIMR-1 localization and establishes SIMR foci as a new germ granule among other known phase separated condensates in the perinuclear space of the C. elegans germline. This work also develops a unique application of proximity-dependent labeling in conjunction with the auxin-inducible degradation system by which to identify previously undiscovered direct interactors of SIMR-1, as well as discover additional SIMR foci components.

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Asset Metadata

Creator Manage, Kevin Indika (author)

Core Title SIMR-1 facilitates robust silencing of piRNA target loci in the C. elegans germline

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Degree Conferral Date 2021-12

Publication Date 09/15/2021

Defense Date 08/19/2021

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

Tag C. elegans,OAI-PMH Harvest,piRNA,RNAi,SIMR foci,SIMR-1,siRNA

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Phillips, Carolyn (committee chair), Benayoun, Berenice (committee member), Curran, Sean (committee member), Michael, Matt (committee member)

Creator Email kmanage2@gmail.com,manage@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-oUC15918309

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