1. Chromosomes and Gene Expression
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A tudor domain protein, SIMR-1, promotes siRNA production at piRNA-targeted mRNAs in C. elegans

  1. Kevin I Manage
  2. Alicia K Rogers
  3. Dylan C Wallis
  4. Celja J Uebel
  5. Dorian C Anderson
  6. Dieu An H Nguyen
  7. Katerina Arca
  8. Kristen C Brown
  9. Ricardo J Cordeiro Rodrigues
  10. Bruno FM de Albuquerque
  11. René F Ketting
  12. Taiowa A Montgomery
  13. Carolyn Marie Phillips  Is a corresponding author
  1. Department of Biological Sciences, University of Southern California, United States
  2. Department of Biology, Colorado State University, United States
  3. Cell and Molecular Biology Program, Colorado State University, United States
  4. Biology of Non-coding RNA Group, Institute of Molecular Biology, Germany
  5. International PhD Programme on Gene Regulation, Epigenetics, and Genome Stability, Germany
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Cite this article as: eLife 2020;9:e56731 doi: 10.7554/eLife.56731

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.

eLife digest

In the biological world, a process known as RNA interference helps cells to switch genes on and off and to defend themselves against harmful genetic material. This mechanism works by deactivating RNA sequences, the molecular templates cells can use to create proteins.

Overall, RNA interference relies on the cell creating small RNA molecules that can target and inhibit the harmful RNA sequences that need to be silenced. More precisely, in round worms such as Caenorhabditis elegans, RNA interference happens in two steps. First, primary small RNAs identify the target sequences, which are then combatted by newly synthetised, secondary small RNAs. A number of proteins are also involved in both steps of the process.

RNA interference is particularly important to preserve fertility, guarding sex cells against ‘rogue’ segments of genetic information that could be passed on to the next generation. In future sex cells, the proteins involved in RNA interference cluster together, forming a structure called a germ granule. Yet, little is known about the roles and identity of these proteins.

To fill this knowledge gap, Manage et al. focused on the second stage of the RNA interference pathway in the germ granules of C. elegans, examining the molecules that physically interact with a key protein. This work revealed a new protein called SIMR-1.

Looking into the role of SIMR-1 showed that the protein is required to amplify secondary small RNAs, but not to identify target sequences. However, it only promotes the creation of secondary small RNAs if a specific subtype of primary small RNAs have recognized the target RNAs for silencing.

Further experiments also showed that within the germ granule, SIMR-1 is present in a separate substructure different from any compartment previously identified. This suggests that each substep of the RNA interference process takes place at a different location in the granule.

In both C. elegans and humans, disruptions in the RNA interference pathway can lead to conditions such as cancer or infertility. Dissecting the roles of the proteins involved in this process in roundworms may help to better grasp how this process unfolds in mammals, and how it could be corrected in the case of disease.

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; 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 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; Liu et al., 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 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 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; 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). 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, 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). 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; 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.

Figure 1 with 1 supplement see all
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 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, 5 μm. (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, 5 μm. (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.

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 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.

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; Zhang et al., 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; Liu et al., 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.

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).

Figure 2 with 1 supplement see all
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.

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., 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 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.

Figure 3 with 1 supplement see all
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 hr 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 for more details regarding statistical analysis.

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 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 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; Gartner et al., 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 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-1, is sufficient to desilence a sensitized piRNA sensor strain, but cannot reactivate a piRNA sensor silenced by RNAe.

Figure 4 with 1 supplement see all
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, 50 μm. (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, 5 μm. (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.

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 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.

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 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). 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). 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). These data implicate SIMR-1 in the production or maintenance of small RNAs at many C. elegans genes.

Figure 5 with 1 supplement see all
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 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 for more details regarding statistical analysis.

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 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-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 (Belicard 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; Wu et al., 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). 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), 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 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 (Figure 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 and Emmons, 1989; Yuan et al., 1991; Collins and Anderson, 1994; Rezsohazy, 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 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.

Figure 6 with 1 supplement see all
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 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 for more details regarding statistical analysis.

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 (Figures 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 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-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). 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 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 and I, 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 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). 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 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.

Figure 7 with 1 supplement see all
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 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 5 μm.

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 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 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 Figures 4B, 5C–D and 6B–E), 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 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 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). 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 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.

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; Yang et al., 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). 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 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).

Plasmid and strain construction

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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 (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 ~1 kb 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 ~500 bp - 1 kb 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. 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. For the introduction of the R159C mutation in SIMR-1::gfp::3xFLAG, we used an oligo repair template and RNA guide (Supplementary file 7).

CRISPR injections were performed according to published protocols (Dickinson et al., 2013; Dickinson et al., 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-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

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~500,000 synchronized N2 (wild-type) or USC717 (mut-16(cmp3[mut-16::gfp::3xFLAG + loxP])) adult C. elegans (~68 hr 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 and FLAG immunoprecipitation was performed at 4°C for 2 hr 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

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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 hr 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 scored per genotype and condition. Information regarding statistical analysis provided in Supplementary file 8.

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, x2-x12+ y2-y12+ z2-z12. 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

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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 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

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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

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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 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.

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.

Reestablishing WAGO-class 22G-siRNA production

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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 individual plates as L4 larvae and scored 2–3 days later for presence or absence of progeny.

Western blots

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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

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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 7.5 μg of each RNA sample, extracted from whole animals, to a final volume of 100 μL. Samples were incubated at 65°C for 2 min then incubated on ice. The Dynabeads mRNA Purification Kit (ThermoFisher 61006) was used according to the manufacturer’s protocol. 20 μL of Dynabeads was used for each sample. 100 ng 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

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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; Liao et al., 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: -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, 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.

Accession numbers

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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).

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Decision letter

  1. Oliver Hobert
    Reviewing Editor; Howard Hughes Medical Institute, Columbia University, United States
  2. James L Manley
    Senior Editor; Columbia University, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

The paper provides interesting new insights into small RNA biology and we thank you for sending this story to eLife.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "A Tudor domain protein, SIMR-1, promotes siRNA production at piRNA-targeted mRNAs in C. elegans" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that we cannot presently accept the manuscript for publication in eLife.

As you will see in the detailed comments by the reviewers below, we appreciate the many interesting insights that your manuscript describes. However, there was an overall concern that, while interesting, the many observations described do not coalesce into a coherent picture at present. Several of the results presented were considered too preliminary and not fully flashed out. You will see that the reviewers propose a number of additional experiments. However, if you are able to successfully address the reviewers’ concerns, eLife may be interested in looking at a new version of this manuscript. Since the required revisions are quite substantial, such a manuscript would need to be considered a new submission.

Reviewer #1:In C. elegans, small RNAs antisense to mRNAs regulate gene expression. The small RNAs are synthesized by RNA-dependent RNA polymerases (RdRPs) that accumulate in perinuclear condensates called mutator foci. How specific mRNAs are chosen for small RNA synthesis is not known, but genomically-encoded piRNAs appear to be involved for at least a subset of loci including transposable elements.

The manuscript by Manage et al., reports on a new component of the C. elegans small RNA synthesis machinery. SIMR-1 contains a TUDOR-like domain, a domain often found in Argonaute-interacting proteins. Proteomic analyses of SIMR-1 indicate that SIMR-1 interacts with MUT-16, other Mutator foci proteins, and WAGO-1, an Argonaute that also interacts with MUT-16. Like Mutator foci proteins, SIMR-1 is required for the synthesis and/or maintenance of small RNAs targeting many loci (>600), including ~100 loci also targeted by piRNAs. SIMR-1 localizes to a new granule adjacent to Mutator foci. The authors speculate that SIMR-1 may function in the "handoff" of mRNAs targeted by piRNAs to mutator foci. This conclusion, however, does not seem well supported by the data since SIMR-1 mutants affect many more genes beyond those targeted by piRNAs (point #1). SIMR-1 interacts with the Argonaute WAGO-1, but a functional connection between SIMR-1 and WAGO-1 was not explored (point #5).

1) The authors do a comprehensive analysis of small RNA defects in SIMR-1 mutants raised at different temperatures and identify nearly ~1000 affected loci. Consistent with SIMR-1 interacting with the Mutator scaffold MUT-16, >90% of the genes affected in simr-1 mutants are also affected in mut-16 mutants. These genes include genes targeted by piRNAs, but the latter represent only a minority (<20%) of genes affected in simr-1 mutants, so it is not clear why the authors focus so much on those genes.

2) 'simr-1 mutants fail to produce high levels of siRNAs from the majority of piRNA-target loci' Figure 4C seems to contradict this statement since at 20°C only <50% of piRNA-target loci are affected. Also, what criteria were used to define "piRNA-targeted loci"? New analyses by the Mello and Lee labs have identified piRNA binding sites in 100s of mRNAs targeted by PRG-1/piRNA complexes. What happens to those loci in simr-1 mutants? Are the small RNAs completely gone or reduced? Are small RNAs nearest piRNA-binding sites most or least affected? is there a specific pattern of loss?

3) The authors argue that the defect must be "downstream of the piRNAs" because piRNAs levels overall are not affected in SIMR-1 mutants. They don't report however the levels of the specific piRNAs that target the ~100 piRNA-targeted genes affected in the simr-1 mutant. The levels of PRG-1 protein (piRNA associated Argonaute) also are not reported.

4) Do the small RNA changes observed in simr-1 mutants correlate with changes in mRNA levels?

5) SIMR-1 interacts with the Argonaute WAGO-1, but the authors do not explore potential functional connections between SIMR-1 and WAGO-1 – why? Is there a good overlap between WAGO-1-dependent loci and SIMR-1-dependent loci?

6) Figure 6: Systematic quantification of relative granule positions shown is missing. See Wan et al., 2018, for examples of quantification metrics. The authors' findings suggest that they are 4 different types of perinuclear condensates in germ cells – are they all present in a 1:1:1:1 proportion? Is every P granule next to a Z granule next to a SIMR-1 granule next to a mutator focus or does the order and composition vary?

Reviewer #2:In this manuscript, Manage et al. have identified a previously uncharacterized protein, called SIMR-1, which interacts with a component of mutator foci, MUT-16, and participates in the accumulation of a subset of endo-siRNAs antisense to piRNA and mutator target genes. They also show that simr-1 mutant displays a temperature-dependent progressive loss of fertility similarly to mutation in downstream components of the piRNA pathway. Interestingly, they have shown that SIMR-1 has a perinuclear localization in a region distinct from p-granule, z-granule or mutator foci, and that its localization is independent from mutator foci. Moreover, they have identified by sequence alignment an extended Tudor domain protein which might potentially interact with methylated PIWI protein. Based on these results, they propose that SIMR-1 constitutes a distinct germ-granule that acts downstream of PIWI and piRNAs to transport mRNAs targeted by piRNAs from p-granule to mutator foci (the granule involved in the amplification of endo-siRNAs). Overall, the distinct perinuclear localization of this protein is exciting, however there are no experiments that substantiate the claim that this protein has a role in transporting mRNAs form piRNA pathway to downstream mutator foci nor that the putative Tudor domain is functional. I also express some concerns about the methods, data analysis, and interpretation of some key results as listed below.

A) The authors performed a MUT-16 IPs coupled with mass spectrometry to identify novel interacting factors of mutator complex. My first concern is related to the fact that the authors did not specify how many IP replicates they have used and which statistical methods they have performed to identify specific interactors in MUT-16 IPs versus control IPs. It appears that they have considered interacting proteins that have peptides only in MUT-16 GFP and FLAG IPs and not in the control. This approach can limit the number of interacting proteins identifiable in the IPs. Also, the number of peptides they have identified for SIMR-1 is extremely low (2 peptides in FLAG IPs and 1 peptide in GFP IPs) rising doubt that SIMR-1 is a real interactor of MUT-16. Because of the lack of statistical tools used to identify interacting proteins I suggest them to perform Co-IP experiments to confirm this interaction. It is also not known whether this interaction is direct or mediated by RNA, and an experiment such as Co-IP can help to distinguish the nature of their interactions. Also, in Figure 1A they have included the proteins RSD-2, WAGO-1, and MATH-33, which have been identified following another (not clear) approach (their peptides are not uniquely present in MUT-16 IPs). Therefore, it should be removed from this table. Again, if they want to take in consideration such cases they should perform a quantitative proteomic approach, using more replicates and statistical tools.

B) There are no functional data or structural information to asses that the conserved residues contained in SIMR-1 constitute a Tudor domain capable of recognizing a methylated substrate such as PIWI. Moreover, SIMR-1 IP and mass spec experiments failed to identify PRG-1/PIWI protein. The major interactor identified by mass spec is instead RSD-2, which they show to colocalize in the same perinuclear foci as SIMR-1, although it appears to have a different function (RSD-2 is involved in RNAi and not 22G-RNA amplification from piRNA targets and SIMR-1 is not involved in RNAi but in the accumulation of 22G-RNAs from piRNA targets). Therefore, the distinct localization of SIMR-1 per se does not implicate that SIMR-1 mediates the handoff of mRNAs from the piRNA pathway to the downstream Mutator components. Also, the peptide counts of the putative interactors of SIMR-1 shown in Supplementary Figure 6 are extremely low, a factor that rises concerns on the quality of the IP/mass spec experiments. Again, it is not specified how many replicates have been performed. Therefore, some of these interactions, such as MUT-16/SIMR-1 should be confirmed by Co-IP.

C) The authors performed a small RNA-seq analysis to check whether specific population of small RNAs are affected in simr-1 mutant. They have performed profiles of small RNAs in different conditions (normal temperature of grow or 25°C at several generations), perhaps to correlate gene expression changes with the temperature-dependent progressive loss of fertility phenotype. However, the lack of corresponding RNA-seq profiles does not allow to investigate the effect of temperature on gene expression and how it correlates with the progressive sterility phenotype. Moreover, the most dramatic changes in small RNAs in SIMR-1 mutant occur at 25°C after just one generation. Thus, there is no correlation between progressive loss of fertility and changes in small RNAs. I wonder then if the results across generation of SIMR-1 mutant grown at 25°C are relevant for the current manuscript at all. Therefore, the only conclusion from all the analyses performed is that SIMR-1 is somehow participating in the accumulation of a subset of mut-16 and piRNA targets. The effect of the temperature on small RNA biogenesis is instead not clear.

D) In addition, the small RNA analysis they have performed is not very compelling. For instance, in some cases is not clear if the criteria they have used to define genes with decrease or increase small RNAs. Also, the methods don't clearly show which small RNAs they are considering (22G-RNAs, 26G-RNAs, sense, antisense, broad range of small RNAs 21-26nt, etc.). Because these are small RNAs cloned from total RNAs and not from AGO IPs, the authors cannot be sure that the increases or decreases of small RNAs observed belong to a specific AGO pathway (it is known that mutation in some small RNA biogenesis factor causes misrouting of small RNAs into AGOs). In general, it appears that mutation in SIMR-1 causes a general downregulation (not completely loss) of small RNAs, which are considered to be mut-16 dependent (including some previously defined piRNA targets). Among these affected genes, the piRNA targets appear more affected than the whole mut-16 targets.

E) The authors claim that SIMR-1 acts downstream of the piRNA pathway, however they haven't performed any experiment to prove that piRNA silencing is impaired in simr-1 mutant. They should use the canonical piRNA sensor strain (Ashe et al., 2012), and check whether the silencing of the piRNA sensor reporter is impaired in simr-1 mutant. This experiment goes well with the other assays they have used to show that RNAi is not affected in simr-1 mutant.

Specific comments:

1) Add in Figure 2C the statistical significance of those changes.

2) Add in Figure 2D genes that are statistically significant up-regulated and down-regulated (or >2-fold change). They can label the significant dots with different colours as well. Same in 2E, if any.

3) In Figure 4C the total number of genes belonging to piRNA, ERGO-1, and mut-16 is changing in the different Venn diagram. This is not clear. Shouldn't be constant?

4) Subsection “simr-1 mutants have increased levels of germ cell apoptosis”, the paragraph title is misleading. The level of germline apoptosis in SIMR-1 is similar to WT at 20°C and lower than WT at 25°C (first generation). It is slightly higher than the corresponding WT at generation 2, 10, 11 and not changing at generation 4,7. Importantly, the WT at 25°C (1 generation) has the highest level of germline apoptosis compared to all the other data points and yet is still fertile. Therefore, the reported level of germline apoptosis is not correlated with the sterility phenotype. This is why I disagree with the Authors' interpretation of the results shown in Figure 3E and the conclusion they wrote in the subsection “simr-1 mutants have increased levels of germ cell apoptosis”.

5) The first paragraph of the subsection “SIMR-1 is required for small RNA production at piRNA target genes”, contains results that are not shown in any figures. Please, provide a figure that reports these results. Also, it is not written which criteria they have used to identify changes in small RNA populations (fold changes, p-value?), nor it is known from the Materials and methods or text which small RNAs they are considering (22G-RNAs, 26G-RNAs, sense, antisense, broad range of small RNAs 21-26nt, etc.). Also, do the small RNA-depleted genes at 25°C (which are not depleted in subsequent generations) reacquire the small RNAs in progressive generations?

6) The results from the analyses shown in the subsection “simr-1 mutants have increased levels of siRNAs mapping to histone genes”, are also not clear and maybe not relevant for the current manuscript. The results described lack a figure reference. Are they shown in Supplementary Figure 5B? Also, in Supplementary Figure 5B they only show depleted siRNAs and not enriched siRNAs. Also, they only consider lists of genes depleted of 22G-RNAs. Maybe if they consider log2(fold changes) they will find more insightful results. In regard to the small RNAs targeting histone mRNAs they have cited a submitted paper (not sure if it is possible to cite the journal where the paper is submitted) and conclude this might be relevant. Mutation in MUT-16, which is linked to piRNA pathway, do not show the same histone small RNAs. Moreover, histone small RNAs are present at all temperature, therefore also do not correlate with the progressive loss of fertility in SIMR-1 mutant. Again, I am not sure how this paragraph fits with the whole manuscript.

7) In the Discussion, the authors claim that their work "provides insight into how mRNAs move between the piRNA pathway and the mutator complex and suggests a role for multiple perinuclear condensates to promote piRNA-mediated mRNA silencing". This has not been demonstrated. There are no experiments showing that the mRNAs targeted by piRNAs move through perinuclear SIMR-1 condensates to Mutator foci, nor that SIMR-1 is required for piRNA silencing. The reduction in a subset of piRNA targets might be an indirect consequence of SIMR-1 function, which still remains unknown. Thus, I suggest to remove this sentence. Same for sentences in the last paragraph of the subsection “RSD-2 and SIMR-1 promote the interaction between distinct primary and secondary siRNA pathways”.

8) In the Materials and methods section the authors should add the detailed protocol used for Immunoprecipitation and mass spec (buffer used etc.). The referenced Uebel et al. do not contain such information. Also, the raw proteomic data has not been released. Moreover, they should add a full description of the small RNA analysis performed as mentioned above.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "A Tudor domain protein, SIMR-1, promotes siRNA production at piRNA-targeted mRNAs in C. elegans" for further consideration by eLife. Your revised article has been evaluated by James Manley as the Senior Editor, and Oliver Hobert as the Reviewing Editor.

The reviewers appreciate the revisions. Before we can accept the manuscript, we ask you to please consider some of the editorial revisions that both reviewers have raised and that appear reasonable. No further experimentation is required. We do not expect the revised paper to go back to the reviewers.

Reviewer #1:The authors have addressed our concerns with new experiments that support the main thesis of the paper: SIMR-1 functions in the piRNA silencing pathway.

Reviewer #2:In this revised manuscript, Manage et al. have addressed most of the major points I have raised in my review. Specifically, they have included data showing that SIMR-1 protein with mutation in a critical residue for the structure of the extended Tudor domain fails to localize to perinuclear granule and affects the fertility of the animal at 25°C, similar to the loss of function allele. Moreover, they have placed the reduction in small RNAs upon mutation of SIMR-1 in the context of piRNA pathway. Indeed, the mutant protein fails to silence a piRNA reporter transgene, in accordance to what I previously asked to show. They have also changed the text according to my suggestion and added more statistical tests (where it was possible) upon my request. However, I do have some additional comments that the authors should address:

1) Even if the Authors have removed all statements that "SIMR-1 "hands-off" RNAs from the piRNA to mutator pathway" in the main text, the passage in the Abstract that says "However, how mRNAs first recognized by the piRNA pathway are then handed off to the mutator pathway is unclear. Here, we identify the Tudor domain protein, SIMR-1, as acting at this step between piRNA production and mutator complex-dependent siRNA biogenesis" is still confusing, suggesting that SIMR-1 acts at that step.

2) The Authors have shown that there is a gradual reduction in small RNAs in simr-1 mutant compared to WT across generations at 25°C in the new Figure 5—figure supplement 1D. However, the heat map shown in Figure 5C suggests that the reduction in small RNAs (even at generation 10) in simr-1 mutant is milder compared to prg-1 mutant. Therefore, I recommend to include in Figure 5—figure supplement 1D data from prg-1 mutant at 20°C and to calculate the significant changes in siRNAs between prg-1 and simr-1 mutants. If the small RNA depletion in prg-1 mutant is more severe than in simr-1 mutant (as it appears from Figure 5C), then the Authors should reconsider their claim that SIMR-1 act downstream the piRNA pathway to produce siRNAs in mutator foci. SIMR-1 might cooperate to accumulate small RNAs downstream of piRNAs together with other pathways or proteins (acting together or independent of SIMR-1 in piRNA-dependent small RNA accumulation). Also, the effect on small RNA accumulation can be still indirectly linked to piRNA-dependent small RNA biogenesis, since no interaction between SIMR-1 and PIWI or MUT-16 has been shown by Co-IP. Therefore, these alternative possibilities should be discussed in the text.

3) The increased siRNAs in simr-1 mutant also appear to be less severe than piRNA mutant (Figure 5C). Similarly, the changes in mRNAs (including histone mRNAs) might be less severe that piRNA mutant. In my opinion this can be an important determinant in explaining the different phenotype between piRNA and simr-1 mutants.

4) In the paragraph now called "simr-1 Mrt phenotype is associated with increased levels of germ cell apoptosis" the Authors speculate that the significant increase in apoptotic cells at later generation in simr-1 mutant compared to the relative WT is correlated to sterility. As I have already mentioned in my first review, although WT animals at 25°C have levels of apoptotic cells similar to those of later generation of sterile simr-1 mutant at 25°C, yet they are fertile. Thus, the number of apoptotic cells detected with this assay in simr-1 mutant do not suggest that they contribute to sterility. As a comparison, mutants of H3K9me (met-2, set-25) show 4-fold increases in apoptotic cells at 25°C compared to WT (in Zeller et al., 2016, Figure 2B). Therefore, in my opinion this data is inconclusive and can induce to misinterpretation. This is why, if the Authors are willing to show this result, they should comment on the fact that WT animals with similar number of apoptotic cells are fertile and that the number of apoptotic cells detected in simr-1 mutant might not contribute to the sterility.

5) In the Introduction, the Authors wrote: "Here we identify a protein required to mediate the interaction between PRG-1 and piRNAs in P granules with siRNA amplification in Mutator foci." The interaction between these factors has not been confirmed by Co-IP or other experiments, thus the statement needs to be changed.

6) In the Introduction, the Authors claim that "this work identifies one of the first factors to act downstream of PRG-1 to mediate the production of secondary siRNAs". This is not accurate because many factors involved in the accumulation of piRNA-dependent small RNAs have been identified thus far. Also, the result presented pointed out that simr-1 might directly or indirectly participate in the accumulation of those siRNAs (see also comment 2).

https://doi.org/10.7554/eLife.56731.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Reviewer #1:

[…] The manuscript by Manage et al., reports on a new component of the C. elegans small RNA synthesis machinery. SIMR-1 contains a TUDOR-like domain, a domain often found in Argonaute-interacting proteins. Proteomic analyses of SIMR-1 indicate that SIMR-1 interacts with MUT-16, other Mutator foci proteins, and WAGO-1, an Argonaute that also interacts with MUT-16. Like Mutator foci proteins, SIMR-1 is required for the synthesis and/or maintenance of small RNAs targeting many loci (>600), including ~100 loci also targeted by piRNAs. SIMR-1 localizes to a new granule adjacent to Mutator foci. The authors speculate that SIMR-1 may function in the "handoff" of mRNAs targeted by piRNAs to mutator foci. This conclusion, however, does not seem well supported by the data since SIMR-1 mutants affect many more genes beyond those targeted by piRNAs (point #1). SIMR-1 interacts with the Argonaute WAGO-1, but a functional connection between SIMR-1 and WAGO-1 was not explored (point #5).

We have removed all statements that SIMR-1 “hands-off” RNAs from the piRNA to mutator pathway and have instead indicated that SIMR-1 acts at a step between piRNA production and mutator-complex-dependent siRNA biogenesis. To provide additional evidence that SIMR-1 acts in the piRNA pathway and not the downstream mutator pathway, we have included data showing that simr-1 mutants, like mutants in the piRNA pathway, de-silence a sensitized piRNA sensor but not a piRNA sensor silenced by RNAe or an ERGO-1 dependent siRNA sensor (Figure 4A and Figure 4—figure supplement 1A-B). Furthermore, SIMR-1 is required to promote fertility after re-establishing production of WAGO-class 22G-siRNAs, a phenotype also associated only with mutants in the piRNA pathway (Figure 4B).

1) The authors do a comprehensive analysis of small RNA defects in SIMR-1 mutants raised at different temperatures and identify nearly ~1000 affected loci. Consistent with SIMR-1 interacting with the Mutator scaffold MUT-16, >90% of the genes affected in simr-1 mutants are also affected in mut-16 mutants. These genes include genes targeted by piRNAs, but the latter represent only a minority (<20%) of genes affected in simr-1 mutants, so it is not clear why the authors focus so much on those genes.

In the first version of this manuscript, we used Supplementary Table 2 from Lee et al., 2012, to define piRNA targets. This list contains approximately 300 genes for which the authors observed 8-fold enrichment of 22G-siRNAs in FLAG::PRG-1 compared to prg-1 mutants and is likely not a comprehensive list of prg-1-dependent 22G-siRNA target loci. Therefore, we generated new small RNA libraries from wild-type and prg-1 mutants at 20°C and defined prg-1-dependent siRNA target genes as those with at least 2-fold reduction in total small RNAs and at least 10RPM in wild-type or mutant libraries (the same criteria by which we defined simr-1-dependent siRNA target genes). Using these new libraries, we generated a more comprehensive list of 1758 prg-1-dependent siRNA target genes. With this larger list, now 686 (out of 817) or 84% of simr-1-dependent siRNA target genes overlap with prg-1-dependent siRNA target genes at 20°C (Figure 5D). At 25°C, 693 (out of 1258) or 55% of simr-1-dependent siRNA target genes overlap with prg-1-dependent siRNA target genes (Figure 5—figure supplement 1C). It is important to note that in the latter case the simr-1 libraries, collected at 25°C, are being compared to prg-1 libraries collected at 20°C and this difference in temperature may account for the lower overlap between simr-1 and prg-1-dependent siRNA target genes. At both temperatures, the majority of simr-1 targets (94% at 20°C and 80% at 25°C) still overlap with mut-16-dependent siRNA target genes (Figure 5D and Figure 5—figure supplement 1C).

We have also, since the first submission, obtained two alleles of simr-1 out of a screen to de-silence a piRNA sensor strain (Figure 4A) and demonstrated that SIMR-1 is required to promote fertility after re-establishing WAGO-class 22G-siRNA production, a phenotype previously observed only with mutants in known piRNA pathway factors prg-1, pid-1, and henn-1 (Figure 4B), providing further evidence that SIMR-1 is important for production of siRNAs at piRNA-target loci.

2) 'simr-1 mutants fail to produce high levels of siRNAs from the majority of piRNA-target loci' Figure 4C seems to contradict this statement since at 20°C only <50% of piRNA-target loci are affected. Also, what criteria were used to define "piRNA-targeted loci"? New analyses by the Mello and Lee labs have identified piRNA binding sites in 100s of mRNAs targeted by PRG-1/piRNA complexes. What happens to those loci in simr-1 mutants? Are the small RNAs completely gone or reduced? Are small RNAs nearest piRNA-binding sites most or least affected? is there a specific pattern of loss?

This sentence was changed to say that simr-1 mutants fail to produce high levels of siRNAs at many piRNA-target loci. See previous response for more detail on how we defined piRNA-target loci (those with at least 2-fold reduction in total small RNAs in prg-1 mutants and at least 10RPM in wild-type or prg-1 libraries). While there is a significant overlap between the genes we defined as simr-1-dependent siRNA loci and prg-1-dependent siRNA loci there are also many prg-1-dependent siRNA loci that do not meet the criteria that we used to define simr-1-dependent siRNA loci. Note, however, that many of these had reduced siRNAs in the simr-1 mutant but did not meet the 2-fold cutoff we used (see boxplot in Figure 5B where more than 75% of piRNA target genes have reduced siRNAs in a simr-1 mutant).

Regarding piRNA binding sites, we decided to focus specifically on piRNAs whose expression changes in a simr-1 mutant (Figure 5F and Supplementary file 4). We identified 77 piRNAs (less than 1% of all piRNAs) that are at least 2-fold down-regulated in a simr-1 mutant. Using piRTarBase (which compiles piRNA binding sites identified by the Mello and Lee labs) we identified 37 mRNAs that are predicted targets of these piRNAs by the piRScan relaxed targeting rules (Lee lab) and the CLASH data (Mello lab). We chose to examine sites that were in common between both of these methods because Wu et al., 2019, demonstrate that only when they examined the piRNA target sites in common between these two methods was it predictive of an increase in mRNA expression and a reduction in 22G levels in a prg-1 mutant; genes identified by either method alone were not significantly predictive. Of these 37 mRNAs that are predicted targets of piRNAs affected in a simr-1 mutant, only five have reduced levels of siRNAs in a simr-1 mutant (see Supplementary file 4). These data indicate that the simr-1-depleted piRNAs are not a major driver of siRNA depletion in simr-1 mutants.

3) The authors argue that the defect must be "downstream of the piRNAs" because piRNAs levels overall are not affected in SIMR-1 mutants. They don't report however the levels of the specific piRNAs that target the ~100 piRNA-targeted genes affected in the simr-1 mutant. The levels of PRG-1 protein (piRNA associated Argonaute) also are not reported.

See previous response. We first identified all piRNAs whose expression is reduced by 2-fold in simr-1 mutants (77 total piRNAs) and then demonstrate that the predicted target mRNAs for these piRNAs are, overall, not depleted of small RNAs in the simr-1 mutant. In total, only five of the 817 genes reduced of small RNAs by at least 2-fold are predicted to be direct targets of these 77 “simr-1-dependent” piRNAs. We have additionally added a western blot for PRG-1 showing that the levels of PRG-1 are not affected by the simr-1 mutant (Figure 7—figure supplement 1C).

4) Do the small RNA changes observed in simr-1 mutants correlate with changes in mRNA levels?

The majority of the genes that are depleted of small RNAs in simr-1 are not affected at the mRNA level. In contrast, the genes that are enriched for small RNAs tend to have reduced expression at the mRNA level (Figure 6H). The failure of the genes depleted of small RNAs to increase in expression at the mRNA level is consistent with the recent observations by Reed et al., 2019, looking at all mutator targets and Barucci et al., 2020, looking at piRNA targets. In both cases, the majority of genes that were depleted of small RNAs failed to increase in mRNA expression. This data indicates that there may be additional layers of gene regulation that maintain silencing of these genes in the absence of siRNAs.

5) SIMR-1 interacts with the Argonaute WAGO-1, but the authors do not explore potential functional connections between SIMR-1 and WAGO-1 – why? Is there a good overlap between WAGO-1-dependent loci and SIMR-1-dependent loci?

MUT-16 interacts via immunoprecipitation and yeast two-hybrid with WAGO-1, but we have no evidence that SIMR-1 interacts with WAGO-1. Therefore, we did not follow up on this possibility.

6) Figure 6: Systematic quantification of relative granule positions shown is missing. See Wan et al., 2018, for examples of quantification metrics. The authors' findings suggest that they are 4 different types of perinuclear condensates in germ cells – are they all present in a 1:1:1:1 proportion? Is every P granule next to a Z granule next to a SIMR-1 granule next to a mutator focus or does the order and composition vary?

We have added quantification showing the percentage of SIMR-1 foci that are associated with P granules, Z granules, and Mutator foci. Additionally, we have included quantification of the distances between fluorescence centers for each of these foci relative to SIMR-1 foci (Figure 7B) and pointed out that in some cases we can observe multiple SIMR-1 foci interacting with another type of granule (see Figure 7A, arrow in inset for SIMR-1 and ZNFX-1 localization). These data show that while SIMR-1 is usually associated with the other three granules, it is not necessarily in a 1:1:1:1 proportion. We have also noticed that not all P granules are associated with SIMR/Mutator/Z granules. We do not know at this time why only some P granules are associated with these other granules however we are quantifying and exploring further in another manuscript.

Reviewer #2:

In this manuscript, Manage et al. have identified a previously uncharacterized protein, called SIMR-1, which interacts with a component of mutator foci, MUT-16, and participates in the accumulation of a subset of endo-siRNAs antisense to piRNA and mutator target genes. They also show that simr-1 mutant displays a temperature-dependent progressive loss of fertility similarly to mutation in downstream components of the piRNA pathway. Interestingly, they have shown that SIMR-1 has a perinuclear localization in a region distinct from p-granule, z-granule or mutator foci, and that its localization is independent from mutator foci. Moreover, they have identified by sequence alignment an extended Tudor domain protein which might potentially interact with methylated PIWI protein. Based on these results, they propose that SIMR-1 constitutes a distinct germ-granule that acts downstream of PIWI and piRNAs to transport mRNAs targeted by piRNAs from p-granule to mutator foci (the granule involved in the amplification of endo-siRNAs). Overall, the distinct perinuclear localization of this protein is exciting, however there are no experiments that substantiate the claim that this protein has a role in transporting mRNAs form piRNA pathway to downstream mutator foci nor that the putative Tudor domain is functional. I also express some concerns about the methods, data analysis, and interpretation of some key results as listed below.

We have removed all statements that SIMR-1 transports RNAs from the piRNA to mutator pathway and have instead indicated that SIMR-1 acts at a step between piRNA production and mutator-complex-dependent siRNA biogenesis. We have also added data demonstrating that the Tudor domain is functional (see response to point “B” below). Finally, we have addressed all specific comments about data analysis and would be happy to provide any other information, if requested by the reviewer.

A) The authors performed a MUT-16 IPs coupled with mass spectrometry to identify novel interacting factors of mutator complex. My first concern is related to the fact that the authors did not specify how many IP replicates they have used and which statistical methods they have performed to identify specific interactors in MUT-16 IPs versus control IPs. It appears that they have considered interacting proteins that have peptides only in MUT-16 GFP and FLAG IPs and not in the control. This approach can limit the number of interacting proteins identifiable in the IPs. Also, the number of peptides they have identified for SIMR-1 is extremely low (2 peptides in FLAG IPs and 1 peptide in GFP IPs) rising doubt that SIMR-1 is a real interactor of MUT-16. Because of the lack of statistical tools used to identify interacting proteins I suggest them to perform Co-IP experiments to confirm this interaction. It is also not known whether this interaction is direct or mediated by RNA, and an experiment such as Co-IP can help to distinguish the nature of their interactions. Also, in Figure 1A they have included the proteins RSD-2, WAGO-1, and MATH-33, which have been identified following another (not clear) approach (their peptides are not uniquely present in MUT-16 IPs). Therefore, it should be removed from this table. Again, if they want to take in consideration such cases they should perform a quantitative proteomic approach, using more replicates and statistical tools.

Only a single replicate was performed for each immunoprecipitation (MUT-16-FLAG, MUT-16-GFP, control-FLAG, control GFP) so we did not perform statistical analysis. The approach was treated as a screen, to identify candidate genes for further exploration, rather than a comprehensive list of true MUT-16 interactors. We have removed RSD-2, WAGO-1, and MATH-33 from the table in Figure 1, as requested. Because we did perform further analysis with these proteins (localization and small RNA seq) that appear in other figures, they were instead included in Figure 1—figure supplement 1, along with the peptide counts in the control strains to make clear that these three proteins were identified at much lower coverage in the control FLAG IP.

We have attempted to confirm the SIMR-1-MUT-16 co-IPs to demonstrate that this is a true interaction, however our results were inconclusive. We believe that this result suggests that the interaction between MUT-16 and SIMR-1 may be weak or transient, which is not surprising given their localization to distinct compartments and the fact that SIMR-1 is not required for exogenous RNAi or small RNA production at many mutator target genes.

B) There are no functional data or structural information to asses that the conserved residues contained in SIMR-1 constitute a Tudor domain capable of recognizing a methylated substrate such as PIWI. Moreover, SIMR-1 IP and mass spec experiments failed to identify PRG-1/PIWI protein. The major interactor identified by mass spec is instead RSD-2, which they show to colocalize in the same perinuclear foci as SIMR-1, although it appears to have a different function (RSD-2 is involved in RNAi and not 22G-RNA amplification from piRNA targets and SIMR-1 is not involved in RNAi but in the accumulation of 22G-RNAs from piRNA targets). Therefore, the distinct localization of SIMR-1 per se does not implicate that SIMR-1 mediates the handoff of mRNAs from the piRNA pathway to the downstream Mutator components. Also, the peptide counts of the putative interactors of SIMR-1 shown in Figure 6—figure supplement 1 are extremely low, a factor that rises concerns on the quality of the IP/mass spec experiments. Again, it is not specified how many replicates have been performed. Therefore, some of these interactions, such as MUT-16/SIMR-1 should be confirmed by Co-IP.

To demonstrate that the Tudor domain of SIMR-1 is functional, we made a single point mutation in a critical residue for the structure of the extended Tudor domain and found that SIMR-1 no longer localizes to perinuclear granules (Figure 4C). Furthermore, these mutant animals have transgenerational fertility defects at elevated temperature similar to the simr-1 deletion allele (Figure 4D). This same allele was also identified in an unbiased screen for de-silencing of a piRNA sensor (Figure 4A). While this data does not prove that the SIMR-1 Tudor domain can recognize methylated peptides, it does support the identification of the Tudor domain and indicate its critical role in SIMR-1 function.

We have removed the SIMR-1 mass-spec experiment due to the low peptide counts and lack of replicates. We attempted to replicate the MUT-16-SIMR-1 interaction by co-IP, but as described in the previous point, the results were inconclusive. Nonetheless, SIMR-1 clearly has a role in the production of siRNAs from piRNA targets based on its phenotypes. We plan to revisit the direct interactors of SIMR-1 in future work.

C) The authors performed a small RNA-seq analysis to check whether specific population of small RNAs are affected in simr-1 mutant. They have performed profiles of small RNAs in different conditions (normal temperature of grow or 25°C at several generations), perhaps to correlate gene expression changes with the temperature-dependent progressive loss of fertility phenotype. However, the lack of corresponding RNA-seq profiles does not allow to investigate the effect of temperature on gene expression and how it correlates with the progressive sterility phenotype. Moreover, the most dramatic changes in small RNAs in SIMR-1 mutant occur at 25°C after just one generation. Thus, there is no correlation between progressive loss of fertility and changes in small RNAs. I wonder then if the results across generation of SIMR-1 mutant grown at 25°C are relevant for the current manuscript at all. Therefore, the only conclusion from all the analyses performed is that SIMR-1 is somehow participating in the accumulation of a subset of mut-16 and piRNA targets. The effect of the temperature on small RNA biogenesis is instead not clear.

We have added a panel showing a reduction in small RNAs mapping to piRNA target genes in simr-1 mutants over the course of 10 generations at 25°C (Figure 5—figure supplement 1D). Additionally, we have added mRNA-seq data to the manuscript, in part, to address whether we can correlate any gene expression changes to the progressive sterility phenotype. We find that some mutator targets, PRG-1 targets, and spermatogenesis genes are up-regulated exclusively at late (sterile or nearly sterile) generations, however, similar classes of genes are also up-regulated at 20°C (Figure 6G). Thus, it is difficult to attribute the sterility phenotype to reduced or increased expression of a specific gene class. We do observe defects in histone mRNA expression (Figure 6I and Figure 6—figure supplement 1G) which Barucci et al., 2020) propose leads to sterility in prg-1 animals; however, in simr-1 mutants the reduction in histone mRNA expression is observed even at 20°C, which suggests that it cannot be the sole cause of the fertility defect. We therefore agree with the reviewer that much of the changes in small RNA expression occur even at 20°C or after a single generation at 25°C – we hypothesize that this loss of piRNA and mutator-target small RNAs is sensitizing the animals to fertility defects at elevated temperature but this may not be a direct effect.

D) In addition, the small RNA analysis they have performed is not very compelling. For instance, in some cases is not clear if the criteria they have used to define genes with decrease or increase small RNAs. Also, the methods don't clearly show which small RNAs they are considering (22G-RNAs, 26G-RNAs, sense, antisense, broad range of small RNAs 21-26nt, etc.). Because these are small RNAs cloned from total RNAs and not from AGO IPs, the authors cannot be sure that the increases or decreases of small RNAs observed belong to a specific AGO pathway (it is known that mutation in some small RNA biogenesis factor causes misrouting of small RNAs into AGOs). In general, it appears that mutation in SIMR-1 causes a general downregulation (not completely loss) of small RNAs, which are considered to be mut-16 dependent (including some previously defined piRNA targets). Among these affected genes, the piRNA targets appear more affected than the whole mut-16 targets.

We have clarified the figures and figure legends to indicate what classes of small RNAs we are examining. In general, we are looking at total small RNAs mapping to a particular gene class and have not restricted it by size, 5’ nucleotide, or strandedness. For piRNAs, we looked only at small RNAs mapping to annotated piRNA loci. We do not argue that these small RNAs are binding particular Argonaute proteins, only that they map to known mutator, piRNA, CSR-1, etc target genes. Finally, using our new prg-1 mutant small RNA data, we show that the majority of simr-1 targets are also piRNA targets, in addition to being mutator targets (Figure 5D).

E) The authors claim that SIMR-1 acts downstream of the piRNA pathway, however they haven't performed any experiment to prove that piRNA silencing is impaired in simr-1 mutant. They should use the canonical piRNA sensor strain (Ashe et al., 2012), and check whether the silencing of the piRNA sensor reporter is impaired in simr-1 mutant. This experiment goes well with the other assays they have used to show that RNAi is not affected in simr-1 mutant.

We have now added the isolation of two alleles of simr-1 from a piRNA sensor screen (first described in de Albuquerque, Genes Dev 2014) to demonstrate that simr-1 can de-silence a sensitized piRNA sensor (Figure 4A) but not a piRNA sensor silenced by RNAe (Figure 4—figure supplement 1A). Furthermore, we have demonstrated that SIMR-1 is required to promote fertility after re-establishing WAGO-class 22G-siRNA production, a phenotype previously observed only with mutants in known piRNA pathway factors prg-1, pid-1, and henn-1 (Figure 4B), providing further evidence that SIMR-1 is important for production of siRNAs at piRNA-target loci.

Specific comments:

1) Add in Figure 2C the statistical significance of those changes.

The libraries in Figure 2C were considered part of the screen to identify new small RNA factors and were not performed with replicates, so statistics was not included. All subsequent small RNA and mRNA seq focused on simr-1 (Figures 5-6 and Figure 5—figure supplement 1 and Figure 6—figure supplement 1) were performed with multiple replicate libraries and include statistics.

2) Add in Figure 2D genes that are statistically significant up-regulated and down-regulated (or >2-fold change). They can label the significant dots with different colours as well. Same in 2E, if any.

Colors have been changed to highlight genes with log2(fold change) in small RNA abundance greater or less than 1.

3) In Figure 4C the total number of genes belonging to piRNA, ERGO-1, and mut-16 is changing in the different Venn diagram. This is not clear. Shouldn't be constant?

These particular Venn diagrams have been replaced, however in the original figure, overlaps of less than 10 genes were not shown on the diagram for simplicity.

4) Subsection “simr-1 mutants have increased levels of germ cell apoptosis”, the paragraph title is misleading. The level of germline apoptosis in SIMR-1 is similar to WT at 20°C and lower than WT at 25°C (first generation). It is slightly higher than the corresponding WT at generation 2, 10, 11 and not changing at generation 4,7. Importantly, the WT at 25°C (1 generation) has the highest level of germline apoptosis compared to all the other data points and yet is still fertile. Therefore, the reported level of germline apoptosis is not correlated with the sterility phenotype. This is why I disagree with the Authors' interpretation of the results shown in Figure 3E and the conclusion they wrote in the subsection “simr-1 mutants have increased levels of germ cell apoptosis”.

We have changed the title of this section to indicate that the increase in apoptosis is specifically associated with the Mrt phenotype in generations 10 and 11. We agree that it is strange that apoptosis increases so significantly in the first generation at 25°C, however we interpret this to be a response to the heat stress. By generation 2 at 25°C, the animals have acclimated to the elevated temperature and apoptosis is reduced. Only in the late generation 25°C animals do we observe apoptosis elevated in the simr-1 mutant compared to wild-type. While we are not arguing that the apoptosis is causative of the sterility, we do believe that the increase in apoptosis in the late generations, which is far more significant than any differences observed in earlier generations, is correlated with reduced fertility in these animals.

5) The first paragraph of the subsection “SIMR-1 is required for small RNA production at piRNA target genes”, contains results that are not shown in any figures. Please, provide a figure that report these results. Also, it is not written which criteria they have used to identify changes in small RNA populations (fold changes, p-value?), nor it is known from the Materials and methods or text which small RNAs they are considering (22G-RNAs, 26G-RNAs, sense, antisense, broad range of small RNAs 21-26nt, etc.). Also, do the small RNA-depleted genes at 25°C (which are not depleted in subsequent generations) reacquire the small RNAs in progressive generations?

We have added tables to indicate the total number of genes with changes in small RNAs or mRNAs in each genotype and condition (Figure 5A and Figure 6F). Details regarding criteria used to define genes with small RNA and mRNA changes and types of small RNAs being analyzed has been added to appropriate figure legends and Materials and methods.

For the genes that lose small RNAs in simr-1 mutants, the majority are found to lose small RNAs across multiple conditions (temperature or generation). For the genes that lose small RNAs in simr-1 mutants only in the first generation at 25°C, but not at 20°C or later generations at 25°C (~300 genes in total), some of these genes have increased small RNAs in the wild-type strain only in this condition and are lost in subsequent generations while others lose small RNAs that are then regained at later generations. In the cases of genes that appear to gain small RNAs in simr-1 mutants only in the first generation at 25°C, but not at 20°C or later generations at 25°C, many of these genes actually have reduced small RNAs in the wild-type strain at this generation that are recovered at later generations. In fact, there are many genes that gain and lose small RNAs in the wild-type strain at elevated temperature and across generations, but this analysis seemed beyond the scope of the current paper so we did not include the data.

6) The results from the analyses shown in the subsection “simr-1 mutants have increased levels of siRNAs mapping to histone genes”, are also not clear and maybe not relevant for the current manuscript. The results described lack a figure reference. Are they shown in Supplementary Figure 5B? Also, in Supplementary Figure 5B they only show depleted siRNAs and not enriched siRNAs. Also, they only consider lists of genes depleted of 22G-RNAs. Maybe if they consider log2(fold changes) they will find more insightful results. In regard to the small RNAs targeting histone mRNAs they have cited a submitted paper (not sure if it is possible to cite the journal where the paper is submitted) and conclude this might be relevant. Mutation in MUT-16, which is linked to piRNA pathway, do not show the same histone small RNAs. Moreover, histone small RNAs are present at all temperature, therefore also do not correlate with the progressive loss of fertility in SIMR-1 mutant. Again, I am not sure how this paragraph fits with the whole manuscript.

We believe that the histone data supports our argument that simr-1 acts in the piRNA pathway (and not the mutator pathway). The paper referenced in the original submission of this manuscript is now published in Nucleic Acids Research (Reed et al., 2019). An additional paper has also recently been published on this topic (Barucci et al., 2020), both demonstrating that increased small RNAs mapping to histone genes and reduced histone mRNA expression is a phenotype associated with the piRNA pathway. Both papers also demonstrate that these small RNAs mapping to histones are dependent on the mutator complex for their biogenesis, thus if they are present, the mutator complex must be functional. Therefore, these data places simr-1 in the piRNA pathway, upstream of the mutator complex-dependent biogenesis of these histone small RNAs.

The sentences which were lacking a figure reference have been removed. Supplementary Figure 5B has been removed and replaced with Figure 6B-C, which shows genes enriched for small RNAs in prg-1, mut-16, and simr-1 mutants, and the correlation of those enriched for small RNAs in prg-1 and simr-1 mutants with histone genes. All of the lists of genes enriched for small RNAs are described in Figure 5A and Supplementary file 3.

7) In the Discussion, the authors claim that their work "provides insight into how mRNAs move between the piRNA pathway and the mutator complex and suggests a role for multiple perinuclear condensates to promote piRNA-mediated mRNA silencing". This has not been demonstrated. There are no experiments showing that the mRNAs targeted by piRNAs move through perinuclear SIMR-1 condensates to Mutator foci, nor that SIMR-1 is required for piRNA silencing. The reduction in a subset of piRNA targets might be an indirect consequence of SIMR-1 function, which still remains unknown. Thus, I suggest to remove this sentence. Same for sentences in the last paragraph of the subsection “RSD-2 and SIMR-1 promote the interaction between distinct primary and secondary siRNA pathways”.

Both sentences have been edited to remove language indicating trafficking or handover between the piRNA pathway and the mutator complex, and simply state that SIMR-1 acts downstream of piRNA biogenesis and upstream of mutator complex-dependent siRNA amplification.

8) In the Materials and methods section the authors should add the detailed protocol used for Immunoprecipitation and mass spec (buffer used etc.). The referenced Uebel et al. do not contain such information. Also, the raw proteomic data has not been released. Moreover, they should add a full description of the small RNA analysis performed as mentioned above.

The IP/mass spec section of the Materials and methods has been updated to include buffers used and the Bioinformatics Analysis section has been modified to include more detail about the packages used and the analysis performed. A table of all mass spec data has been included as Supplementary file 2. We would be happy to provide any additional details, if requested by the reviewer.

[Editors’ note: what follows is the authors’ response to the second round of review.]

Reviewer #2:

[…] 1) Even if the Authors have removed all statements that "SIMR-1 "hands-off" RNAs from the piRNA to mutator pathway" in the main text, the passage in the Abstract that says "However, how mRNAs first recognized by the piRNA pathway are then handed off to the mutator pathway is unclear. Here, we identify the Tudor domain protein, SIMR-1, as acting at this step between piRNA production and mutator complex-dependent siRNA biogenesis" is still confusing, suggesting that SIMR-1 acts at that step.

We have edited the Abstract to eliminate the suggestion that a “hand-off” is occurring.

2) The Authors have shown that there is a gradual reduction in small RNAs in simr-1 mutant compared to WT across generations at 25°C in the new Figure 5—figure supplement 1D. However, the heat map shown in figure 5C suggests that the reduction in small RNAs (even at generation 10) in simr-1 mutant is milder compared to prg-1 mutant. Therefore, I recommend to include in Figure 5—figure supplement 1D data from prg-1 mutant at 20°C and to calculate the significant changes in siRNAs between prg-1 and simr-1 mutants. If the small RNA depletion in prg-1 mutant is more severe than in simr-1 mutant (as it appears from Figure 5C), then the Authors should reconsider their claim that SIMR-1 act downstream the piRNA pathway to produce siRNAs in mutator foci. SIMR-1 might cooperate to accumulate small RNAs downstream of piRNAs together with other pathways or proteins (acting together or independent of SIMR-1 in piRNA-dependent small RNA accumulation). Also, the effect on small RNA accumulation can be still indirectly linked to piRNA-dependent small RNA biogenesis, since no interaction between SIMR-1 and PIWI or MUT-16 has been shown by Co-IP. Therefore, these alternative possibilities should be discussed in the text.

We added a line in Figure 5—figure supplement 1D below which all piRNA targets fall (since the list of piRNA-target genes was defined as those reduced by 2-fold in a prg-1 mutant). We agree that the reviewer makes a good point that piRNA-target genes are reduced of siRNAs in prg-1 more severely than simr-1, even at late generations. Thus, we have adjusted the text to reflect that sterility is unlikely to be caused by this loss progressive loss of siRNAs in simr-1, as prg-1 was fertile at the time it had an even greater loss of siRNAs to these genes. We have additionally modified the first paragraph of the Discussion to point out that the simr-1 phenotypes are weaker than those of prg-1, 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.

3) The increased siRNAs in simr-1 mutant also appear to be less severe than piRNA mutant (Figure 5C). Similarly, the changes in mRNAs (including histone mRNAs) might be less severe that piRNA mutant. In my opinion this can be an important determinant in explaining the different phenotype between piRNA and simr-1 mutants.

As mentioned in previous comment, we have edited the first paragraph of the Discussion to discuss the difference in phenotypes between simr-1 and prg-1 and offer possible reasons for this.

4) In the paragraph now called "simr-1 Mrt phenotype is associated with increased levels of germ cell apoptosis" the Authors speculate that the significant increase in apoptotic cells at later generation in simr-1 mutant compared to the relative WT is correlated to sterility. As I have already mentioned in my first review, although WT animals at 25°C have levels of apoptotic cells similar to those of later generation of sterile simr-1 mutant at 25°C, yet they are fertile. Thus, the number of apoptotic cells detected with this assay in simr-1 mutant do not suggest that they contribute to sterility. As a comparison, mutants of H3K9me (met-2, set-25) show 4-fold increases in apoptotic cells at 25°C compared to WT (in Zeller et al., 2016, Figure 2B). Therefore, in my opinion this data is inconclusive and can induce to misinterpretation. This is why, if the Authors are willing to show this result, they should comment on the fact that WT animals with similar number of apoptotic cells are fertile and that the number of apoptotic cells detected in simr-1 mutant might not contribute to the sterility.

An additional sentence has been added to this section of the Results to make it clear that high levels of apoptosis are not always correlated to sterility.

5) In the Introduction, the Authors wrote: "Here we identify a protein required to mediate the interaction between PRG-1 and piRNAs in P granules with siRNA amplification in Mutator foci." The interaction between these factors has not been confirmed by Co-IP or other experiments, thus the statement needs to be changed.

We have edited this sentence so as not to imply a direct physical interaction between SIMR-1 and these other pathways.

6) In the Introduction, the Authors claim that "this work identifies one of the first factors to act downstream of PRG-1 to mediate the production of secondary siRNAs". This is not accurate because many factors involved in the accumulation of piRNA-dependent small RNAs have been identified thus far. Also, the result presented pointed out that simr-1 might directly or indirectly participate in the accumulation of those siRNAs (see also comment 2).

We have edited this sentence so as not to indicate that SIMR-1 is the first factor identified with this role.

https://doi.org/10.7554/eLife.56731.sa2

Article and author information

Author details

  1. Kevin I Manage

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Conceptualization, Formal analysis, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1992-0782
  2. Alicia K Rogers

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5525-6095
  3. Dylan C Wallis

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-2375-5062
  4. Celja J Uebel

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Funding acquisition, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0362-1238
  5. Dorian C Anderson

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  6. Dieu An H Nguyen

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Resources, Funding acquisition
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4927-7329
  7. Katerina Arca

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  8. Kristen C Brown

    1. Department of Biology, Colorado State University, Fort Collins, United States
    2. Cell and Molecular Biology Program, Colorado State University, Fort Collins, United States
    Contribution
    Formal analysis
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5495-3371
  9. Ricardo J Cordeiro Rodrigues

    1. Biology of Non-coding RNA Group, Institute of Molecular Biology, Mainz, Germany
    2. International PhD Programme on Gene Regulation, Epigenetics, and Genome Stability, Mainz, Germany
    Contribution
    Resources
    Competing interests
    No competing interests declared
  10. Bruno FM de Albuquerque

    Biology of Non-coding RNA Group, Institute of Molecular Biology, Mainz, Germany
    Contribution
    Resources
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8483-6822
  11. René F Ketting

    Biology of Non-coding RNA Group, Institute of Molecular Biology, Mainz, Germany
    Contribution
    Resources, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6161-5621
  12. Taiowa A Montgomery

    Department of Biology, Colorado State University, Fort Collins, United States
    Contribution
    Formal analysis, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7857-3253
  13. Carolyn Marie Phillips

    Department of Biological Sciences, University of Southern California, Los Angeles, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft, Writing - review and editing
    For correspondence
    cphil@usc.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6228-6468

Funding

National Cancer Institute (K22 CA177897)

  • Carolyn Marie Phillips

National Institute of General Medical Sciences (R35 GM119656)

  • Carolyn Marie Phillips

National Institute of General Medical Sciences (T32 GM118289)

  • Dieu An H Nguyen

National Institute of General Medical Sciences (R35 GM119775)

  • Taiowa A Montgomery

Deutsche Forschungsgemeinschaft (KE1888/1-1)

  • Rene F Ketting

Deutsche Forschungsgemeinschaft (KE1888/1-2)

  • Rene F Ketting

Pew Charitable Trusts (Pew Scholar in the Biomedical Sciences)

  • Carolyn Marie Phillips

National Science Foundation (DGE 1418060)

  • Celja J Uebel

University of Southern California (Dornsife College Chemistry-Biology Interface trainee)

  • Celja J Uebel

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

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.

Senior Editor

  1. James L Manley, Columbia University, United States

Reviewing Editor

  1. Oliver Hobert, Howard Hughes Medical Institute, Columbia University, United States

Publication history

  1. Received: March 8, 2020
  2. Accepted: April 24, 2020
  3. Accepted Manuscript published: April 27, 2020 (version 1)
  4. Version of Record published: May 28, 2020 (version 2)

Copyright

© 2020, Manage et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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