A convenient method for the delivery of extracellular dsRNA into C. elegans at various times during larval development is the expression of dsRNA in the bacteria that the animals ingest as food (Timmons and Fire, 1998). To determine when ingested dsRNA can enter the germline and cause silencing, we exposed developing animals with a ubiquitously expressed protein (GTBP-1) tagged with GFP to bacteria that express gfp-dsRNA. Silencing was detectable within the germline from the second larval stage (L2) onwards (Figure 1A, Figure 1—figure supplement 1A), but either exposure to ingested dsRNA beyond the fourth larval stage (L4) (Figure 1B) or injection of dsRNA into the 1-day-old adult germline (Figure 1—figure supplement 1B) was required to observe silencing in the germline of 3-day-old adults. Combined with the need for exposure to dsRNA after the L4 stage (Marré et al., 2016; Wang and Hunter, 2017) for silencing in progeny, even for just 24 hr (Figure 1—figure supplement 1C), these observations suggest that detectable silencing within the germline is possible during early development without initiating heritable RNA silencing. One possible explanation for this observation could be that both RNAs derived from the imported dsRNA and downstream silencing signals are continually diluted by the proliferation of germ cells. Heritable silencing by dsRNA ingested from the L4 stage to the first day of adulthood likely relies on entry of dsRNA into the proximal germline because silencing of a somatic gene in progeny after parental ingestion of dsRNA during this period required RME-2 (Figure 1—figure supplement 1C), which is enriched in the proximal germline (Figure 1—figure supplement 1D; Grant and Hirsh, 1999), and some gtbp-1::gfp animals exposed to gfp-dsRNA up to the first day of adulthood showed more silencing in the proximal germline (Figure 1—figure supplement 1E).

Thus, these results reveal three periods of germline development that can be broadly distinguished based on the response to ingested dsRNA: (1) from the first larval to the fourth larval stage when exposure to dsRNA does not result in maximal silencing within the germline in adults (Figure 1B); (2) from the fourth larval stage to early adulthood when entry of dsRNA primarily occurs in the proximal germline through RME-2 (Figure 1—figure supplement 1C and E); and (3) later adulthood when germline silencing by ingested dsRNA is maximal (Figure 1B) and ingested dsRNA can effectively silence progeny independent of RME-2 (Figure 1—figure supplement 1C; Wang and Hunter, 2017).

Another approach for delivering extracellular dsRNA into the germline that better mimics dsRNA transport between cells is the secretion of dsRNA from neurons (Devanapally et al., 2015). However, the extent of such secretion throughout development is unpredictable. To modulate the secretion of dsRNA from somatic cells into parental circulation during development, we adapted an optogenetic approach for damaging somatic cells (Xu and Chisholm, 2016). Specifically, we generated animals that express the mini singlet oxygen generator (miniSOG) protein in neurons and exposed them to blue light. While animals expressing miniSOG from a single-copy transgene did not show an appreciable defect when compared with wild-type animals, those expressing miniSOG from a multi-copy transgene were paralyzed (Figure 2—figure supplement 1A and B, top) and had visibly damaged neurons (Figure 2—figure supplement 1B, bottom). Using this system, we induced oxidative damage in the neurons of animals that expressed dsRNA under the control of a neuronal promoter and evaluated silencing of target genes with matching sequence expressed in other tissues (Figure 2A). By exposing animals to blue light for 60 min at different times during development (Figure 2—figure supplement 1C), we observed SID-1-dependent enhancement in the silencing of the hypodermal gene bli-1 at the adult stage by neuronal bli-1-dsRNA, with maximal silencing when oxidative damage occurred during mid-to-late larval development (Figure 2—figure supplement 1D, light exposure every 6 hr from 42 to 66 hr post L4-stage of parent; Figure 2—figure supplement 1E, ~two-fold increase from 14.9% to 29.1% in a background with enhanced RNA interference (eri-1(-)) and ~six-fold increase from ~1.6% to ~9.8% in a wild-type background). A similar period of maximal SID-1-dependent enhancement of silencing was also observed when neurons expressing gfp-dsRNA were damaged and silencing of a two-gene operon that expresses two fluorescent proteins, mCherry::H2B and GFP::H2B, in the germline was measured (Figure 2B-D, Figure 2—figure supplement 1F 48–60 hr post L4-stage of parent; sid-1(-) allele (jam80[nonsense]) is depicted in Figure 2—figure supplement 2). While silencing of gfp::h2b was observed throughout the germline, silencing of the other cistron mCherry::h2b was also observed sometimes, albeit restricted to specific regions of the germline. Silencing of mCherry::h2b was most frequent in the proximal germline and was not observed in any other region without silencing in the proximal germline (proximal germline – 57%, distal germline – 47%, sperm – 29%, Figure 2D), likely due to reduction of mCherry::h2b::gfp::h2b pre-mRNA (Guang et al., 2008) in those regions. Consistently, the silencing of both gfp::h2b and mCherry::h2b was eliminated in the absence of the nuclear Argonaute HRDE-1 (Figure 2—figure supplement 1G). The pattern of mCherry::h2b silencing is similar to the spatial pattern observed for the RME-2-dependent entry of dsRNA delivered into parental circulation (Marré et al., 2016) and is consistent with the pattern of target mRNA degradation in the germline by extracellular dsRNA (Ouyang et al., 2022). However, silencing of gfp::h2b in the germline by neuronal dsRNA did not show a detectable dependence on RME-2 (Figure 2—figure supplement 1H - difficulty in obtaining transgenic animals that also lack RME-2 resulted in a low sample size for this experiment).

Thus, by modulating the secretion of dsRNA from somatic cells for the first time, we gained two insights into the intercellular transport of dsRNA: (1) oxidative damage of neurons during particular periods in development increases the amount of dsRNA and/or changes the kinds of dsRNA in circulation either because of specific enhancement of secretion or nonspecific spillage; and (2) there is a preference for silencing by neuronal dsRNA in the proximal germline.

While the characteristics of extracellular dsRNA imported into the germline from ingested bacteria or from neurons are unknown, delivery of chemically defined dsRNA into the extracellular space in C. elegans can be accomplished using microinjection (Fire et al., 1998; Marré et al., 2016). We examined differences, if any, in the entry of in vitro transcribed dsRNA from the extracellular body cavity into the germline during the L4 and adult stages as evidenced by silencing in the progeny of injected animals. Silencing was comparable regardless of whether wild-type or rme-2(-) parents were injected as L4-staged or adult animals (Figure 3A, Figure 3—figure supplement 1A, left; also reported for adults in Wang and Hunter, 2017). However, some dependence on RME-2 for silencing in progeny was discernable when lower concentrations of dsRNA were used (Figure 3—figure supplement 1A, right). This result and previous results demonstrating proximal germline-specific silencing in contexts where silencing is less robust (Figure 2D, Figure 1—figure supplement 1E) are consistent with RME-2-dependent uptake of dsRNA being a route for extracellular dsRNA to enter the germline. The difference in parental RME-2 requirement for silencing in progeny observed for dsRNA ingested (Figure 1—figure supplement 1C) or injected (Figure 3—figure supplement 1A) at the L4 stage could similarly reflect the accumulation of different amounts of dsRNA in parental circulation (e.g. more upon injection than upon ingestion), and/or different kinds of dsRNA (e.g. because of modifications in bacteria or upon transit through the intestine). However, these possibilities could not be easily distinguished because sensitive northern blotting (Choi et al., 2017) revealed that both bacterial and in vitro transcribed dsRNA consist of a complex mix of dsRNAs (Figure 3—figure supplement 1B–1D; consistent with Tabara et al., 2002 and Jain et al., 2020), hereafter called mixed dsRNA.

In contrast, when synthesized gfp-dsRNA of a defined length (50 bp) with a fluorescent label was injected into circulation in adult animals, no entry into the germline was observed in the absence of RME-2 (Marré et al., 2016). We found that silencing of unc-22 in progeny by similarly synthesized but unlabeled 50 bp unc-22-dsRNA with a 5’ OH delivered into parental circulation also showed a strong requirement for RME-2, unlike mixed dsRNA (Figure 3A). Further comparison between the two forms of dsRNA revealed that silencing in progeny by 50 bp dsRNA injected into parental circulation was detectably less efficient in somatic cells (Figure 3A, Figure 3—figure supplement 2A and B, left), even when ~14X more 50 bp dsRNA was delivered into parental circulation (Figure 3—figure supplement 2B, right), and was also less efficient in the germline (Figure 3B, Figure 3—figure supplement 2A and C). Efficient silencing in response to added dsRNA requires nuclear Argonaute proteins: NRDE-3 in somatic cells (Guang et al., 2008) and HRDE-1 in the germline (Buckley et al., 2012). Both 50 bp dsRNA and mixed dsRNA relied on HRDE-1 for silencing within the germline (Figure 3—figure supplement 2A and C) and could silence independent of NRDE-3 in somatic cells (Figure 3—figure supplement 2A and C). In addition to the diversity of RNA lengths observed in mixed dsRNA, another known feature that could distinguish dsRNA transcribed in bacteria or in vitro from synthesized 50 bp dsRNA is the presence of 5’ triphosphates on the transcribed dsRNA species instead of the 5’ OH present in synthesized 50 bp dsRNA. In support of the impact of 5’ phosphates on transport and/or silencing, addition of 5’ monophosphates to synthesized 50 bp dsRNA injected into parental circulation reduced the dependence on RME-2 for silencing in progeny (Figure 3—figure supplement 1E and F), potentially by enhancing the ability of synthesized dsRNA to be imported by other dsRNA importers (e.g. SID-1) in the absence of RME-2. Thus, the requirements for dsRNA entry into the germline and subsequent silencing vary for different lengths and/or chemical forms of dsRNA (see summary in Table 1).

Fluorescently labeled 50 bp dsRNA delivered into parental circulation localized within intestinal cells in progeny (Figure 3C, top left), as has been observed for vitellogenin proteins (Sharrock, 1983) and fluorescent dyes (Bossinger and Schierenberg, 1996). Accumulation of fluorescently labeled dsRNA was also detected at the apical membrane of the intestine, which could reflect exocytosis of dsRNA into the lumen of developing intestinal cells. However, separation of the fluorescent label from dsRNA catalyzed by cellular enzymes cannot be excluded. Therefore, to dissect differences, if any, between the transport of unlabeled short dsRNA (synthesized 50 bp with 5’OH) and mixed dsRNA (mixture transcribed in vitro using ~1 kb DNA template) we injected unc-22-dsRNA into animals with mutations in genes that play roles in the import of dsRNA. We found that maternal SID-1 was required for silencing by short dsRNA in progeny (Figure 3C, bottom, left bars), suggesting that the SID-1-dependent entry of short dsRNA into the cytosol likely occurs in the injected parent or during early development in progeny. Uptake of dsRNA from the intestinal lumen requires SID-2, a transmembrane protein located in the apical membranes of intestinal cells (Winston et al., 2007; McEwan et al., 2012). We found that SID-2 was not required for most silencing in progeny by short or mixed dsRNA injected into parental circulation (Figure 3C, top right and bottom). Exit of dsRNA from intracellular vesicles requires SID-5, a transmembrane protein located in endolysosomal membranes (Hinas et al., 2012). Silencing in wild-type animals was comparable to silencing in sid-5(-) animals (Figure 3C, top right). However, when animals that lacked SID-1 were injected, SID-5 was required in progeny for silencing by mixed dsRNA from parental circulation (Figure 3C, bottom, right bars; as also reported in Wang and Hunter, 2017). Since dsRNA is expected to be present in vesicles upon entry through RME-2 in the absence of SID-1 (Marré et al., 2016; Wang and Hunter, 2017), this observation suggests that SID-5 is required for the release of mixed dsRNA from inherited vesicles in progeny.

In summary, injected extracellular dsRNA can enter the germline in parents and be transmitted to progeny through two routes with different substrate selectivity. One route is preferentially used by short dsRNA and relies on RME-2-mediated endocytosis of dsRNA into oocytes, where early exit from vesicles is required for silencing in progeny as evidenced by the need for maternal SID-1 (Figure 3D, blue). The other route appears to exclude short dsRNA but allows mixed dsRNA entry into the cytosol in the parental germline through SID-1 and exit from inherited vesicles in progeny through a process that requires both zygotic SID-1 and SID-5 (Figure 3D, grey; Wang and Hunter, 2017).

All routes of dsRNA transport deduced using the experimental addition of dsRNA ultimately require SID-1 for entry into the cytosol. The proposed model (Figure 3D) for dsRNA transport into the germline and to progeny suggests that the expression pattern of SID-1 is likely to include the germline. We used Cas9-mediated genome editing to insert a piRNA-resistant mCherry sequence (Zhang et al., 2018; Devanapally et al., 2021) into the sid-1 coding sequence (Figure 4A) to observe the endogenous expression pattern of SID-1::mCherry. This fusion protein was detectably functional in contrast to other attempts at tagging SID-1 (see Technical comments in Materials and methods and Figure 4—figure supplement 1). Fluorescence from SID-1::mCherry was visible in the proximal and distal regions of the adult germline (Figure 4B). Expression also progressively increased during development with tissue-specific enrichment in the developing embryo (Figure 4C, left), becoming ubiquitous in hatched L1 larvae (Figure 4C, middle, and Figure 4D). SID-1::mCherry was not easily detectable in the germline during later larval development (Figure 4C, middle and right). In combination with the expression of RME-2 observed in the proximal germline (Figure 1—figure supplement 1D; Grant and Hirsh, 1999), this expression pattern of SID-1 is consistent with the entry of most dsRNA from circulation of adult animals into the germline, followed by activity of transport mechanisms in developing embryos and early larvae that inherit parental dsRNA.

To determine if acute, induced expression rather than developmental expression of SID-1 can be sufficient for the import of dsRNA into the germline, we engineered the endogenous sid-1 gene to transcribe a fusion transcript with an aptamer-regulated ribozyme (Figure 4—figure supplement 2A, left) that cleaves itself when not bound to tetracycline (Figure 4—figure supplement 2A, right) (based on Wurmthaler et al., 2019). Exposing these animals to tetracycline enabled silencing by dsRNA in somatic tissues (hypodermis: Figure 4—figure supplement 2B, left; body-wall muscles: Figure 4—figure supplement 2B, right), indicative of stabilization of sid-1 mRNA, production of SID-1 protein, and subsequent dsRNA import in somatic cells. However, such tetracycline-induced silencing was not detectable in the germline (Figure 4—figure supplement 2C-F). Yet, similar tagging of the ubiquitously expressed gene gtbp-1::gfp resulted in detectable rescue of expression within the germline by tetracycline (Figure 4—figure supplement 2G). A possible explanation for the poor rescue of SID-1 activity within the germline is that post-transcriptional mechanisms targeting sid-1 mRNA in the germline but not in somatic cells interfere with tetracycline-dependent stabilization of the sid-1 transcript (e.g. piRNA-based regulation of sid-1 mRNA, Ouyang et al., 2019; Dodson and Kennedy, 2019), or that acute stabilization of the sid-1 transcript does not override developmental regulation of SID-1 translation.

Additional attempts to tag the SID-1 protein guided by structure and to modulate sid-1 transcripts guided by post-transcriptional regulatory interactions could improve control of dsRNA transport between cells. Nevertheless, the developmentally regulated expression observed for both SID-1 and RME-2 in the germline is consistent with intergenerational or transgenerational effects of endogenous dsRNA from parental circulation after development of the adult germline.

To understand how the dsRNA importer SID-1 might be used in endogenous gene regulation across generations, we searched for sid-1-dependent changes in gene expression that could be heritable (Figure 5, Figure 2—figure supplement 2 and Figure 5—figure supplement 1). To control for genetic background (see Technical comments in Materials and methods), we used Cas9-mediated genome editing to delete the entire sid-1 coding sequence or introduce a nonsense mutation in cohorts of the same wild-type animals. By comparing polyA+ RNA from this wild type with that of the newly generated sid-1(jam113[deletion]) (Figure 5A and B, Figure 5—figure supplement 1A) or sid-1(jam80[nonsense]) (Figure 5A–C) animals, we found that 26 genes were significantly (q<0.05) misregulated in sid-1(jam113[deletion]) (Figure 5—figure supplement 1B) and 6 in sid-1(jam80[nonsense]) (Figure 5D, top), both including sid-1 (a list of significantly altered genes is in Table 2). The most upregulated gene in sid-1(jam113[deletion]), F14F9.5 (Figure 5—figure supplement 1B), which is located immediately 3’ to sid-1 in the genome, was only misregulated in the deletion mutant and not in the nonsense mutant (Figure 5—figure supplement 1C, left). Both mutants, however, were equally defective for silencing by ingested dsRNA (Figure 5B). This observation suggests that while both mutations result in loss of SID-1 protein, the deletion of sid-1 also changes local regulatory sequences (potentially explaining upregulation of the neighboring gene F14F9.5) and eliminates sid-1 mRNA, which could participate in RNA-based regulatory interactions within the germline (Ouyang et al., 2019; Dodson and Kennedy, 2019). Nevertheless, we could detect two genes that were upregulated in both sid-1(jam113[deletion]) and sid-1(jam80[nonsense]) animals (red in Figure 5D, top, and Figure 5—figure supplement 1B): the identical loci W09B7.2/F07B7.2 (Figure 5—figure supplement 1C, middle), and Y102A5C.36 (Figure 5—figure supplement 1C, right). Intriguingly, another gene cls-3 also changed in both mutants (Table 2) but in different directions (~3.4-fold decrease in the sid-1(jam80[nonsense]) mutant but a ~5.8-fold increase in the sid-1(jam113[deletion]) mutant), suggesting that the direction of change in expression can vary. Conservatively, we began by analyzing only the two genes with mRNA levels that changed in the same direction in both sid-1 mutants. Both W09B7.2/F07B7.2 and Y102A5C.36 have been reported (Reed et al., 2020) to be expressed within the germline (Figure 5—figure supplement 1D, left) and regulated by endogenous small RNAs (Figure 5—figure supplement 1D, middle and right). Spliced mRNA levels measured at a later generation using RT-qPCR demonstrated that both transcripts were upregulated in sid-1(jam80[nonsense]) animals compared to wild-type animals as expected (Figure 5E), but no upregulation was detectable in sid-1(jam113[deletion]) animals (Figure 5—figure supplement 1E). This difference between the two sid-1 mutants could reflect increased variation in expression (as was observed for cls-3 using RNAseq) or could reflect complex effects caused by deletion of sid-1 DNA (e.g. F14F9.5 overexpression, loss of sid-1 mRNA, etc.) that could be independent of SID-1 protein function.

To determine if changes in W09B7.2/F07B7.2 and Y102A5C.36 expression were heritable, we reverted the sid-1 nonsense mutation to wild-type sequence using Cas9-mediated genome editing. This immediately restored silencing by ingested dsRNA (Figure 5B) with concomitant recovery of sid-1 mRNA to wild-type levels (Figure 5E, top). In contrast, changes in both W09B7.2/F07B7.2 and Y102A5C.36 expression persisted (Figure 5D, bottom) even after one year of passaging the reverted animals (sid-1(jam86[revertant]); i.e. after >100 generations, Figure 5E, middle and bottom). Thus, the sid-1-dependent accumulation of mRNA from these two germline genes persisted for many generations, likely through mechanisms that maintain heritable epigenetic changes. We hereafter refer to these sid-1-dependent genes (sdg) that show heritable epigenetic changes in response to temporary loss of SID-1 as sdg-1 (W09B7.2/F07B7.2) and sdg-2 (Y102A5C.36).

To determine if expression of sdg-1 and sdg-2 is regulated by other proteins that play a role in RNA silencing within the germline, we examined 21 published datasets (Buckley et al., 2012; Dodson and Kennedy, 2019; Welker et al., 2007; Spike et al., 2008; Wan et al., 2021; Suen et al., 2020; Wahba et al., 2021; Shen et al., 2018; Goh et al., 2014; Lee et al., 2012; Batista et al., 2008; Kim et al., 2021; Ni et al., 2016) that reported changes that depend on such proteins. For each dataset, we determined if the lists of genes reported as showing significant changes in mutants compared to the respective wild types included sdg-1 and/or sdg-2. This analysis revealed that changes in mRNA and/or antisense small RNAs of sdg-1 were detected in 20 of the 21 datasets while changes in sdg-2 were observed in 9 of 21 (Figure 5F). When detected, changes in sdg-2 were in the same direction as changes in sdg-1, suggestive of similar regulation of both genes.

RNAs transcribed in the germline can be recognized as they exit the nuclear pores by piRNAs bound to the Argonaute PRG-1, which recruits them for regulation by antisense small RNAs called 22G RNA made by proteins organized within perinuclear germ granules (reviewed in Sundby et al., 2021). Interaction with piRNAs was detected for RNA from sid-1, sdg-1, and sdg-2, and the control gene tbb-2 using crosslinking, ligation, and sequencing of hybrids (Shen et al., 2018; Figure 5F), consistent with their germline expression. Depletion of downstream 22G RNAs targeting both sid-1 and sdg-1 was detectable upon loss of the germ granule component MUT-16 (Suen et al., 2020; Figure 5F). Both genes were among the top 500 genes targeted by 22G RNAs bound by the secondary Argonaute HRDE-1/WAGO-9 (Buckley et al., 2012; Figure 5F), suggesting similar downregulation of both genes using 22G RNAs. Furthermore, multiple datasets support downregulation of sdg-1 within the germline by HRDE-1/WAGO-9-bound 22G RNAs in the absence of PRG-1. One, loss of HRDE-1/WAGO-9 increased sdg-1 RNA in whole animals (Ni et al., 2016; Figure 5F) and in dissected gonads (Kim et al., 2021; Figure 5F). Two, loss of PRG-1 decreased sdg-1 RNA (Figure 5F) and increased 22G RNAs that are antisense to sdg-1 (Figure 5F) in dissected gonads (Wahba et al., 2021). Three, although animals that lack PRG-1 become progressively sterile, the increase in sdg-1 22G RNA persisted in near-sterile animals (Figure 5F, near-sterile in Wahba et al., 2021), and this increase was eliminated upon additional loss of HRDE-1/WAGO-9 (Figure 5F, near-sterile in Wahba et al., 2021).

As expected for sid-1-dependent downregulation of sdg-1, multiple datasets support an inverse relationship between the two genes. In animals lacking PRG-1, sid-1 RNA levels increased and sid-1 22G RNAs decreased (Lee et al., 2012; Figure 5F), but both sdg-1 RNA and sdg-2 RNA levels decreased along with an increase in 22G RNAs (Suen et al., 2020; Wahba et al., 2021; Shen et al., 2018; Goh et al., 2014; Lee et al., 2012; Batista et al., 2008; Figure 5F). This inverse relationship between sid-1 and sdg-1 RNA regulation is also observed when many components of germ granules are mutated as indicated by changes in 22G RNA upon loss of the embryonic P granule components MEG-3/-4 (Dodson and Kennedy, 2019; Figure 5F), the PRG-1 interactor DEPS-1 (Suen et al., 2020; Figure 5F), or the Z granule component ZSP-1 (Wan et al., 2021; Figure 5F; also known as PID-2, Placentino et al., 2021).

In addition to the above studies, pioneering studies that used microarrays identified sdg-1 as upregulated in animals lacking the germ granule component DEPS-1 (Spike et al., 2008; Figure 5F) and in animals lacking the dsRNA-binding protein RDE-4 (Figure 5F; second-most upregulated in Welker et al., 2007), which recruits dsRNA imported through SID-1 and other intracellular dsRNA for processing and eventual gene silencing. Animals that lack RDE-4 show a ~47.5-fold increase in sdg-1 RNA (Welker et al., 2007). A reduction in RDE-4 activity could also contribute to the ~11.6-fold increase in sdg-1 RNA seen in deps-1(-) animals because these animals also show a ~3.2-fold decrease in rde-4 RNA (one of 13 downregulated genes in Spike et al., 2008). These observations support the idea that appropriate regulation of sdg-1 RNA requires both piRNA-mediated processes that act via germ granule components such as DEPS-1 and dsRNA-mediated processes that use SID-1 and RDE-4.

In summary, the levels of sdg-1 RNA are detectably regulated by the dsRNA-selective importer SID-1, the dsRNA-binding protein RDE-4, and the piRNA-binding Argonaute PRG-1. Presence of dsRNA-mediated regulation or loss of piRNA-mediated regulation enhances MUT-16-dependent production of secondary small RNAs that bind the secondary Argonaute HRDE-1/WAGO-9. Consistent with downregulation of these sid-1-dependent transcripts by SID-1, disruption of many components of germ granules results in opposite effects on these transcripts and sid-1 RNA. Intriguingly, a search of protein interaction studies revealed that the SDG-1 protein is among the interactors of two germ granule components: PID-2 by immunoprecipitation (Placentino et al., 2021; also known as ZSP-1, Wan et al., 2021) and DEPS-1 by proximity labeling (Price et al., 2021). This interaction raises the possibility that the sdg-1 gene is not only regulated by germ granule components, but also that its protein product associates with one or more components of germ granules.

Given these observations on the regulation of sdg-1 and the potential for the SDG-1 protein to be a regulator of RNA silencing, we focus here on the analysis of this gene. The sdg-1 gene is located within a retrotransposon (Figure 6—figure supplement 1A) that is within a duplicated ~40 kb region and has two recognizable paralogs (Figure 6—figure supplement 1B). However, there are no detectable SDG-1 homologs in species other than C. elegans, suggesting that it is a recently evolved gene. Potentially because of this lack of conservation, AlphaFold is unable to predict a confident structure for SDG-1 (CELE_F07B7.2 on the Database; Varadi et al., 2022). To facilitate analysis of SDG-1 expression, we tagged both loci that encode SDG-1 with mCherry coding sequences lacking piRNA-binding sites (mCherry∆pi; Zhang et al., 2018; Devanapally et al., 2021; Figure 6—figure supplement 1C and D). This tagging resulted in the expression of sdg-1::mCherry∆pi mRNA being ~16-fold higher than sdg-1 mRNA (Figure 6—figure supplement 1E), potentially because of the reduction in the overall density of piRNA-binding sites per transcript, the additional introns included in mCherry∆pi (based on Okkema et al., 1993 and Crane et al., 2019), and/or other unknown factors. Fluorescence from SDG-1::mCherry was observed in the germline of adult animals (Figure 6A). However, animals showed variation in SDG-1::mCherry expression between their two gonad arms (Figure 6A, middle shows bright anterior [20% of animals] and right shows bright posterior [6% of animals]). A contributing feature for the observed stochasticity could be the location of sdg-1 within a duplicated region (Figure 6—figure supplement 1A), as suggested by similar stochastic RNA silencing of multi-copy transgenes but not single-copy transgenes (Le et al., 2016). Despite this variation, unbiased passaging of self-progeny for more than 18 generations continuously preserved SDG-1::mCherry expression in an otherwise wild-type background (Figure 6B). In contrast, mating, which can perturb RNA regulation within the germline in cross progeny (Devanapally et al., 2021), caused dramatic changes in sdg-1 expression that persisted in descendants (Figure 6C). Mating animals that express SDG-1::mCherry with wild-type animals resulted in heritable changes along lineages that express sdg-1::mCherry∆pi mRNA or that express sdg-1 mRNA (Figure 6C and Figure 6—figure supplement 2A). This discovery of mating-induced perturbation in gene expression raises caution in interpreting past studies (summarized in Figure 5F) where changes in multiple independent isolates were not examined. Nevertheless, when we used genetic crosses to determine the impact of mutations on sdg-1 expression, we observed reduced SDG-1::mCherry fluorescence in mutants predicted to have reduced levels of intracellular dsRNA (Figure 6D, sid-2(-), sid-5(-)) or reduced processing of intracellular dsRNA (Figure 6D, eri-1(-)). In contrast, we observed an increase in SDG-1::mCherry fluorescence in animals lacking MUT-16 (Figure 6D). Finally, animals lacking RME-2, which lack the ability to import many maternal factors (e.g. lipids, proteins, RNAs, etc.), also showed an increase in SDG-1::mCherry fluorescence (Figure 6D).

To avoid mating-induced perturbations of RNA regulation within the germline, we used Cas9-mediated genome editing to introduce mutations into animals that express SDG-1::mCherry in an otherwise wild-type background. Use of this approach to mutate a control gene with no known roles in RNA regulation within the germline resulted in similar levels of SDG-1::mCherry fluorescence in multiple isolates of animals with and without the mutation (Figure 6—figure supplement 2B). In contrast, mutating sid-1 using Cas9-mediated genome editing caused a range of expression levels in different isolates when compared with sid-1(+) animals (Figure 6E - 6 isolates lower, 2 isolates comparable, and 1 isolate higher), which differs from the increase in sdg-1 mRNA observed upon SID-1 loss in the single isolate examined earlier (Figure 5). Mutating sid-3 also lowered the levels of SDG-1::mCherry in one isolate, but caused no detectable change in another (Figure 6F). While both isolates with loss of RDE-1 showed lower levels of SDG-1::mCherry, both isolates with loss of the germ granule component DEPS-1 showed higher levels of SDG-1::mCherry (Figure 6F). These experiments also reveal that in most isolates of animals expected to have reduced levels of intracellular dsRNA (sid-3(-), sid-1(-)) or reduced processing of intracellular dsRNA (rde-1(-)), SDG-1::mCherry fluorescence is reduced. In contrast, isolates lacking DEPS-1 showed increased expression of SDG-1::mCherry.

Collectively, the observations on SDG-1 expression using mutants suggest that the uptake and processing of intracellular dsRNA (influenced by SID-1, SID-2, SID-3, SID-5, RDE-1, and ERI-1) and the function of germ granules (influenced by MUT-16 and DEPS-1) are both necessary to maintain intermediate levels of SDG-1 expression across generations. Once the levels of SDG-1::mCherry were reduced upon loss of SID-1, downregulation persisted across generations even after restoration of wild-type SID-1 (Figure 6G), just as the upregulation of untagged sdg-1 mRNA also persisted (Figure 5). Despite >100 generations of such persistent silencing, the expression of SDG-1::mCherry could be restored by mutating deps-1 (Figure 6G), implicating small RNA-based regulation and germ granules in the maintenance of the new epigenetic states established upon loss of SID-1.

Since SDG-1 interacts with regulators of RNA silencing (ZSP-1/PID-2, Placentino et al., 2021) and DEPS-1 (Price et al., 2021), loss of SID-1 could both block the entry of extracellular dsRNA into the cytosol and change intracellular RNA regulation through SID-1-dependent genes such as sdg-1. In support of this possibility, intracellular delivery of dsRNA through injection into the syncytial germline of sid-1(-) animals showed a weak defect in silencing (Figure 2D in Marré et al., 2016 and Figure 1 in Wang and Hunter, 2017). To examine if changes in the levels of SDG-1 alone could account for such a defect in silencing by intracellular dsRNA in the germline, we exposed independently-generated sdg-1 deletion animals and animals that overexpress sdg-1 (i.e. with sdg-1::mCherry∆pi) to dsRNA matching the germline gene pos-1 for ~16 hr. This short exposure to pos-1-dsRNA caused intermediate levels of silencing in wild-type animals and comparable intermediate silencing in sdg-1 mutant and sdg-1 overexpressing animals (Figure 7A), suggesting that changes in sdg-1 levels alone are not sufficient to cause a defect in silencing by dsRNA that is detectable using this assay. Alternatively, the previously described defect in silencing by intracellular dsRNA in sid-1(-) animals (Marré et al., 2016; Wang and Hunter, 2017) could be through the promotion of competing piRNA-mediated gene regulation in the absence of SID-1. This notion that dsRNA-mediated and piRNA-mediated gene regulation compete in the germline is supported by a study that demonstrated that loss of PRG-1 enhances heritable RNA silencing by dsRNA (Shukla et al., 2021). To test if loss of SID-1 or a sid-1-dependent gene enhances piRNA-mediated silencing, we used an experimental system which initiates piRNA-mediated silencing of the two-gene operon described in Figure 2 through mating, independent of externally provided dsRNA (Devanapally et al., 2021). We found that sid-1(-) animals exhibited enhanced mating-induced silencing (Figure 7B, top right:~50% off in sid-1(+) vs 100% off in sid-1(qt9[nonsense])), while animals lacking sdg-1 showed a small reduction in mating-induced silencing that was not statistically significant (Figure 7B, bottom right, ~40% off in sdg-1(+) vs ~30% off in sdg-1(jam232[deletion])). Taken together, these results support the model that loss of SID-1 weakly inhibits silencing by intracellular dsRNA (Marré et al., 2016; Wang and Hunter, 2017) but enhances silencing by piRNAs within the germline.

RNA regulation within the germline using piRNAs relies on phase-separated granules (reviewed in Dodson and Kennedy, 2020). To determine if the previous identification of SDG-1 as a putative interactor of the Z-granule component PID-2 (Placentino et al., 2021)/ZSP-1 (Wan et al., 2021) and potentially of the P-granule-adjacent protein DEPS-1 (Price et al., 2021) could be seen as colocalization with Z-granules in vivo, we examined the localization of SDG-1::mCherry within the cytoplasm at high resolution using AiryScan imaging (Huff et al., 2017). SDG-1::mCherry was enriched in perinuclear foci in many animals (Figure 7C, top; 7 of 9 animals) and these sites of enrichment overlapped with the Z-granule marker GFP::ZNFX-1 (Ishidate et al., 2018; Wan et al., 2018; Figure 7C, bottom; 100% colocalized in 10 of 12 animals with enrichment). However, given the resolution of our imaging, we cannot distinguish between a specific interaction with one of the many Z granule components (Zhao et al., 2024) and an interaction with components of the many other nearby perinuclear granules that continue to be reported (Huang et al., 2024). Therefore, we conservatively conclude that SDG-1::mCherry colocalizes with perinuclear germ granules. Time-course imaging revealed re-localization of SDG-1::mCherry into the nucleus from the cytoplasm in the –1 oocyte, which will be the first to be fertilized (Figure 7D, left and Video 1), along with subsequent exclusion from the nucleus before the maternal and paternal pronuclei meet (Figure 7D, right and Video 2). Such dynamic entry into the nucleus followed by exclusion from the nucleus also occurred during early cell divisions in the developing embryo (Videos 3 and 4). The timing of nuclear entry of SDG-1::mCherry coincides with the nuclear envelope breakdown events that occur during fertilization and early development (Huelgas-Morales and Greenstein, 2018). The sdg-1 coding sequence was required for regulated nuclear entry because deletion of the sdg-1 open reading frame in sdg-1::mCherry∆pi animals resulted in mCherry expression throughout the germline in both the cytoplasm and nuclei (Figure 7E). Nuclear localization of the SDG-1 protein in the –1 oocyte is like that of the essential Argonaute CSR-1b (Charlesworth et al., 2021), thought to play a role in protecting transcripts from silencing. Exposure to ingested dsRNA did not alter the expression of sdg-1p::mCherry∆pi[sdg-1(∆)]::sdg-1 3' UTR (Figure 7F) but loss of rde-4 perturbed expression such that one isolate showed loss of expression while the other showed enhanced expression (Figure 7G). These results suggest that while the sdg-1 open reading frame is required for exclusion from germline nuclei, it is not required for the response of the sdg-1 gene to changes in intracellular dsRNA. Together, these observations on the levels and localizations of SDG-1 raise the possibility that this protein is actively regulated by extracellular and intracellular dsRNA with a role in heritable RNA silencing, potentially with perinuclear germ granules, and additional cell-cycle coupled roles during early development, potentially through nucleocytoplasmic shuttling.

The temporal and/or spatial preferences observed for silencing by both ingested dsRNA (Figure 1) and neuronal dsRNA (Figure 2) could be because of unknown characteristics of the exported neuronal dsRNA or ingested dsRNA expressed from bacteria (e.g. modifications, lengths, structures, etc.) that influence import or subsequent silencing. This hypothesis is supported by the different genetic requirements reported for silencing by neuronal gfp-dsRNA compared to other sources of gfp-dsRNA (Ravikumar et al., 2019). Alternatively, these preferences could reflect universal constraints for silencing using any extracellular dsRNA (e.g. expression patterns of factors that promote the import or processing of dsRNA). Furthermore, for both neuronal and ingested dsRNA it is unknown if additional extracellular molecules impact the uptake of dsRNA into cells. Regardless of the presence of such unknown molecules that could be released from neurons to a different degree upon oxidative damage, the gene regulatory effects of the released neuronal dsRNA provide convenient readouts that can be analyzed to understand neuronal damage and its consequences in animals. Pathologies of the central nervous system in humans, including cancer, stroke, multiple sclerosis, neurodegenerative disease, and brain injury, have been associated with extracellular RNAs detected in circulation (reviewed in Tielking et al., 2019), although their origins and regulatory consequences, if any, remain unknown. While the physiological conditions that promote secretion of dsRNA from C. elegans neurons are not known, the discovery that oxidative damage of neurons can enhance the secretion of dsRNA suggests that disruption of cell structures by oxidative damage (e.g. membrane integrity) or initiation of cellular processes that repair oxidative damage (e.g. through ejection of damaged macromolecules; Melentijevic et al., 2017) also promote the release of dsRNA.

The trafficking of extracellular dsRNA from parent to progeny has spatial specificity, as evidenced by more silencing within the proximal germline (Figure 2), temporal specificity, as evidenced by the need for dsRNA beyond the fourth larval stage (Marré et al., 2016; Wang and Hunter, 2017; Figure 1), and substrate specificity, as evidenced by the differential requirements for 50 bp dsRNA with 5’ OH versus a mix of longer dsRNAs with 5’ triphosphates (Figure 3). The observed difference in the extent of silencing by these dsRNAs delivered into parental circulation cannot be attributed to differential engagement of the nuclear Argonautes HRDE-1 and NRDE-3 in progeny (Figure 3—figure supplement 2), but rather could be the result of differences in the ability of each type of dsRNA to bind to upstream factors in the RNA interference pathway (e.g. RDE-4), differences in the stability of each type of dsRNA, and/or differences in the intergenerational transport of each type of dsRNA. Drivers of temporal specificity during development include changes in the uptake of dsRNA into the intestine, distribution of dsRNA to other tissues from the intestine, import of dsRNA into the germline, and availability of RNA silencing factors within the germline. Proteins mediating dsRNA transport or subsequent silencing also differ in their availability during development and in their affinities for different substrates. For example, SID-1 was not detectable in the developing larval germline but was detected in the adult germline (Figure 4) and has an extracellular domain that binds dsRNA (Li et al., 2015) but could prefer dsRNA molecules with 5’ phosphates and/or long dsRNA. Another consideration for substrate specificity is the composition of ‘mixed dsRNAs’ generated through in vitro transcription versus defined short dsRNAs. In addition to a variety of lengths, mixed dsRNAs could also include a variety of undefined secondary structures. Such structures could recruit or titrate away RNA-binding proteins in addition to the dsRNA structures engaging the canonical RNAi pathway, resulting in mixed mechanisms of silencing. Additionally, although the selectivity uncovered here could apply to all dsRNA delivered into the extracellular space of C. elegans from any source (see Table 1 for constraints on intergenerational dsRNA transport), the chemistry of the delivered dsRNA could be modified by yet unidentified enzymes in vivo to overcome these requirements. Tracking labeled dsRNA with diverse chemistries from parental circulation to progeny could allow correlation of differences observed in progeny silencing to differences in intergenerational trafficking.

The germline is a major site of dsRNA import in C. elegans as evidenced by three key observations: the expression of SID-1 in the germline (Figure 4), heritable misregulation of germline genes in sid-1(-) animals (Figures 5 and 6), and accumulation of fluorescently labeled dsRNA from the extracellular space in the germline (Marré et al., 2016; Wang and Hunter, 2017). As a result, sid-1(-) animals could have a defect in the germline that is detectable only under conditions that promote dsRNA transport (e.g. oxidative damage). Multiple physiological defects in the germline and soma of sid-1(-) animals have been reported but have not been widely reproduced, have only been characterized within single generations, or have not been attributed to any specific sid-1-dependent gene(s). These include defects in animals with some misfolded proteins in the endoplasmic reticulum (Long et al., 2014), in animals exiting the dauer stage (Ow et al., 2018; Kadekar and Roy, 2019), in animals exposed to pathogenic P. aeruginosa (Palominos et al., 2017; Moore et al., 2019; Kaletsky et al., 2020), in animals exposed to an odorant (Posner et al., 2019), in intestinal cells that develop in the presence of a multi-copy transgene (Ohno and Bao, 2022), and in animals that overexpress α-synuclein (Gaeta et al., 2022). RNA-seq experiments in this study and comparisons to those of previous studies suggest that genetic background-dependent changes can obscure genuine sid-1-dependent changes (see Figure 4—figure supplement 1, Figure 5—figure supplement 1 and Technical comments in Materials and methods), raising caution in the interpretation of putative sid-1-dependent defects. Comparing multiple sid-1 mutants generated using genome editing with animals in which the mutated sequence has been reverted to wild-type sequence in the same genetic background could provide a firmer basis for the identification of additional sid-1-dependent processes while avoiding the potential for mating-induced changes in gene regulation (Devanapally et al., 2021) that can result when genetic crosses are used instead of genome editing.

Genes expressed within the germline are likely regulated by positive feedback loops that are required to continually produce factors for maintaining germline immortality and for preserving form and function across generations (Jose, 2020; Chey and Jose, 2022). Thus, germline genes could be particularly vulnerable to heritable epigenetic changes, where deviations in the expression levels of a gene that is regulated by or is part of such feedback loops has the potential to become permanent in descendants. Prior data on sdg-1 mRNA (Figure 5F) and the SDG-1 protein (Placentino et al., 2021; Price et al., 2021) suggested the hypothesis that reduction of sdg-1 RNA via SID-1 alters the amount of SDG-1 protein, which could interact with components of germ granules to mediate RNA regulation within the germline of wild-type animals. Our analysis of sdg-1 expression suggests that it is part of a regulatory architecture that is susceptible to heritable epigenetic changes through the perturbation of RNA regulation (Figures 5—7). Such architectures within the germline could be exploited by ‘selfish’ genetic elements such as retrotransposons to persist across evolution if one of these elements also include genes encoding a regulator. In support of a wider use of such a strategy, a paralog of SDG-1, ZK262.8 (Figure 6—figure supplement 1B), is also encoded by a gene located within a retrotransposon and its loss along with that of the miRNA-associated Argonaute ALG-2 was reported to be synthetic lethal (Tops et al., 2006).

Loss of SID-1 weakly inhibits silencing by intracellular dsRNA (Marré et al., 2016; Wang and Hunter, 2017) but enhances the initiation of mating-induced silencing within the germline (Figure 7B). However, loss of SDG-1 alone does not account for these effects (Figure 7A and B), suggesting that additional SID-1-dependent genes could be involved. Furthermore, the ultimate consequence of SID-1 loss could be different in different isolates as exemplified by the differential changes in cls-3 in sid-1(jam80[nonsense]) and sid-1(jam113[deletion]) animals (Table 2). Indeed, the levels of SDG-1 also vary in different isolates of sid-1(-) animals (Figure 6), suggesting that in the absence of SID-1 the sdg-1 gene becomes prone to heritable epigenetic changes, resulting in either high or low expression states being stabilized in different isolates. Together these observations suggest that SID-1-dependent import of extracellular dsRNA could play a role in buffering against heritable epigenetic changes in the germline. Such buffering could be necessary to maintain levels of gene expression within a particular range for each regulatory context. Given the enrichment of SDG-1 protein with perinuclear germ granules (Figure 7C) and the maintenance of heritable changes in sdg-1::mCherry∆pi expression by a DEPS-1-dependent process (Figure 6G), buffering against changes in gene expression could involve both RNA- and protein-based regulation that tunes the function of perinuclear germ granules. We therefore speculate that one role for extracellular RNAs that enter germ cells in other systems (e.g. tRNA fragments in mammals; Sharma et al., 2016; Chen et al., 2016; Sharma et al., 2018) could be to similarly buffer against heritable changes in gene expression.

All strains (listed in Table 3) were cultured on Nematode Growth Medium (NGM) plates seeded with 100 μl of OP50 E. coli at 20 °C and strains made through mating were generated using standard methods (Brenner, 1974). Oligonucleotides used are in Table 4 (for genotyping sid-1(qt9): P1-P2, ttTi5605: P3-P5, eri-1(mg366): P6-P7, sid-1(tm2700): P8-P10, hrde-1(tm1200): P11-P13, nrde-3(tm1116): P14-P16, and rde-4(ne301): P156-P157). Strains made through mating existing mutant strains and genotyping using the above primers are listed below.

Animals were transformed with plasmids and/or PCR products using microinjection (Mello et al., 1991) to generate extrachromosomal arrays or single-copy transgenes. All plasmids were purified from bacterial culture using QIAprep Spin Miniprep Kit (QIAGEN) and all PCR products were generated with Phusion High-Fidelity DNA Polymerase (New England BioLabs) and purified using NucleoSpin Gel and PCR Clean-up Kit (Macherey-Nagel).

Genome editing was performed by injecting nuclear-localized Cas9 (PNA Bio) preincubated at 37 °C for 10 min with either a single-guide RNA (sgRNA) generated by in vitro transcription (SP6 RNA Polymerase, New England BioLabs) or hybridized crRNA/tracrRNA (IDT), as well as an oligonucleotide or PCR-amplified homology repair template, into the C. elegans distal gonad. Screening for plates with successfully edited F1 animals was performed using either dpy-10 co-CRISPR (Arribere et al., 2014; Paix et al., 2015) or the pRF4 plasmid used as a co-injection marker (Dokshin et al., 2018). All plasmids were purified from bacterial culture using QIAprep Spin Miniprep Kit (QIAGEN) and all PCR products were generated with Phusion High-Fidelity DNA Polymerase (New England BioLabs) and purified using NucleoSpin Gel and PCR Clean-up Kit (Macherey-Nagel). Alleles generated by genome editing are schematized in Figure 4A (sid-1), Figure 5A (sid-1), Figure 1—figure supplement 1D (rme-2), Figure 2—figure supplement 2 (deps-1, mut-16, sid-2, rme-2, sid-1, rde-1, sid-5, and sid-3), Figure 4—figure supplement 2A (sid-1), and Figure 6—figure supplement 1C (W09B7.2/F07B7.2 (sdg-1)), and oligonucleotides used are in Table 4.

All animals and embryos expressing fluorescent reporters were imaged in 10 μl of 3 mM levamisole on a 2% agarose pad using a Nikon AZ100 microscope and Photometrics Cool SNAP HQ² camera. A C-HGFI Intensilight Hg Illuminator was used to excite mCherry (filter cube: 530–560 nm excitation, 570 nm dichroic, and 590–650 nm emission), GFP or other autofluorescent molecules in the green channel (filter cube: 450–490 nm excitation, 495 nm dichroic, and 500–550 nm emission) and autofluorescent molecules in the blue channel (filter cube: 325–375 nm excitation, 400 nm dichroic, 435–485 nm emission). Intensity of GFP and mCherry were quantified in Fiji (Schindelin et al., 2012; NIH) using the methods described below. Representative images were adjusted in Fiji (Schindelin et al., 2012; NIH) and/or Adobe Photoshop to identical levels for presentation (Figure 2B-D—4B and C, 6A and 7E, Figure 1-figure supplement 1E and Figure 4-figure supplement 2G).

After RNA sequencing samples were processed as described above, reads were mapped to the C. elegans genome (Kim et al., 2019) using the command ‘hisat2 -p 8 x Celegans98index –1 cutread1.gz –2 cutread2.gz -S sam1’. The SAM file outputs were then converted to BAM files (Li, 2009) using the command ‘samtools view -b sam1 | samtools sort ->bam1.bam’ and BAM index files were created for visualization using ‘samtools index bam1.bam’. Reads for the sid-1 and F14F9.5 locus, W09B7.2/F07B7.2 locus, and Y102A5C.36 locus were plotted using custom R scripts and axes were normalized for each sample based on its total mapped reads, calculated using the command ‘samtools view -c -F 4 bam1.bam’ (Figure 5—figure supplement 1C).

Datasets in 21 published studies were collected and compared based on the gene names to identify changes in sid-1, sdg-1, sdg-2 and tbb-2 (control), if reported (Figure 5F). After standardizing the names across all datasets, the fold-changes reported, if available, were used to plot a heatmap. Cases where fold-changes were not available were set conservatively as log₂(fold change)=2. The R script used is available at https://github.com/AntonyJose-Lab/Shugarts_et_al_2023 (AntonyJose-Lab, 2023).

Total RNA was extracted using TRIzol (Fisher Scientific) from 200 μl pellets of mixed-stage animals collected from 3 to 6 non-starved but crowded plates in biological triplicate for each strain. The aqueous phase was then washed with an equal amount of chloroform and precipitated overnight on ice with 100 μl of 3 M sodium acetate, 1 ml of 100% ethanol and 10 μg glycogen (Invitrogen). RNA pellets were washed twice with 70% ethanol and resuspended in 22 μl nuclease-free water. RNA samples were then Dnase-treated in Dnase buffer (10 mM Tris-HCl pH 8.5, 2.5 mM MgCl₂, 0.5 mM CaCl₂) with 0.5 U Dnase I (New England BioLabs) at 37 °C for 60 min followed by heat inactivation at 75 °C for 10 min. RNA concentration was measured and 1 μg of total RNA was used as input for reverse transcription using 50 U SuperScript III Reverse Transcriptase (Invitrogen) (+RT) or no reverse transcriptase as a negative control (-RT) (RT primers: tbb-2 (P98), sid-1 (P101), W09B7.2/F07B7.2 (P104), Y102A5C.36 (P107)). For qPCR, each +RT biological replicate was assayed in technical triplicate for each gene target, along with a single -RT sample for each corresponding biological replicate using 2 μl cDNA and a no template control (NTC) with the LightCycler 480 SYBR Green I Master kit (Roche). Ct values were measured with the Bio-Rad C1000 CFX96 Real-Time System and Bio-Rad CFX Software (qPCR primers: tbb-2 (P99 and P100), sid-1 (P102 and P103), W09B7.2/F07B7.2 (P105 and P106), Y102A5C.36 (P108 and P109)). To calculate relative change in mRNA abundance compared to wild type, we calculated log₂(2^{(-(gene Ct – tbb-2 Ct))}) using the median of technical replicates for the biological triplicates of each genotype. Ct values were only used if they were lower than corresponding -RT and NTC Ct values. The median value of wild-type biological replicates was then subtracted from the value for each sample to plot calculated values with respect to wild-type levels (Figure 5E, Figure 5—figure supplement 1E, Figure 6—figure supplements 1E and 2A).

BLAST (NCBI) searches were performed using the W09B7.2/F07B7.2 (SDG-1) amino acid sequence with default parameters and any homologs identified were aligned to SDG-1 using Clustal Omega (Madeira et al., 2019) with default parameters. Alignments produced are shown in Figure 6—figure supplement 1B with residues shared by two proteins (grey highlight) or all three proteins (black highlight) indicated.

The Cer9 retrotransposon containing W09B7.2/F07B7.2 (sdg-1) was annotated using sequence features from UCSC Genome Browser and amino acid sequences obtained from Bowen and McDonald, 1999. The 5’ and 3’ LTR sequences were identified using RepeatMasker and were confirmed to have TC and GA dinucleotides at the beginning and end of each sequence, respectively (Bowen and McDonald, 1999). Amino acid sequences from Bowen and McDonald, 1999 corresponding to gag and pol (PR: protease, RT: reverse transcriptase, RH: RNaseH, IN: integrase) elements of Cer9 were used in tblastn (NCBI) searches to determine their positions in the Cer9 retrotransposon sequence that also contains sdg-1.

Mating-induced silencing was assayed by crossing males with the transgene labeled T (oxSi487) encoding mex-5p::mCherry::h2b::gfp::h2b to hermaphrodites lacking the transgene, both in otherwise wild-type backgrounds or indicated mutant backgrounds. Reciprocal control crosses were performed in parallel where hermaphrodites with T were crossed to males lacking T. Animals were imaged and scored as described for this transgene in the ‘Light-induced damage of neurons’ section.

Temporal selectivity of dsRNA transport was probed using three approaches for delivery of dsRNA (damage-induced release from neurons, ingestion, and injection). Spatial selectivity of dsRNA import and/or subsequent silencing was inferred based on differences in the frequency of patterns of silencing within the germline. Substrate selectivity of dsRNA transport pathways was probed using genetic mutants and dsRNA of different lengths and 5’ chemistry. Diversity of dsRNAs made in bacteria and upon in vitro transcription was visualized using Northern blotting. Analysis of sid-1 mutants generated from the same wild-type cohort and a revertant was used for better control of genetic background, aiding in the identification of sid-1-dependent genes (sdg). Separate measurement of sdg-1 expression in descendants of independently edited isolates, along different lineages after perturbations, and in different gonads within single animals demonstrated stochasticity in gene expression and revealed establishment of different heritable epigenetic states. Co-localization of SDG-1::mCherry in perinuclear foci with the Z-granule marker GFP::ZNFX-1, its reported association with the Z-granule component ZSP-1/PID-2 and DEPS-1, changes in its levels in response to loss of SID-1 or the dsRNA-binding protein RDE-4 and its dynamic nuclear localization similar to CSR-1b was used to propose that SDG-1 plays a role in small RNA regulation while also being modulated by the activity of SID-1 and RDE-4.