LOTR-1 is a Z-granule protein that contains both LOTUS and Tudor domains

Inaugural papers that first described the LOTUS winged-helix domain and its conservation identified C. elegans D1081.7 as a hypothetical orphan protein with a single LOTUS domain. [12,13] More recently, LOTUS domains have been subdivided into extended LOTUS (eLOTUS) domains (like the LOTUS domain in Drosophila’s Oskar), and minimal LOTUS (mLOTUS) domains that lack a conserved α5 helix (like those present in mammalian MARF1) (S1 Fig). [14] The LOTUS domain of D1081.7 described in these inaugural papers corresponds to the eLOTUS domain (aa 33–130), but our analysis uncovered an accompanying mLOTUS domain (aa 180–285) (Fig 1A). LOTUS domains have low sequence similarity and are challenging to identify using sequence analysis alone. The mLOTUS domain of D1081.7 shares sequence identity (21%) to the winged-helix region of Cdt1, a regulator in the DNA replication complex. [73] The predicted 3D structures of D1081.7 LOTUS domains superimpose well with solved structures from other species: eLOTUS domains of D1081.7 and Oskar align with a root-mean-square deviation (rmsd) of 2.8 Å, whereas mLOTUS, which lacks the α5 helix, deviates from both fly and human LOTUS folds by ~3.5 Å rmsd (Fig 1B). These paired LOTUS domains reflect the arrangement of TDRD5 and TDRD7 in mammals and two proteins recently described in C. elegans called MIP-1 and MIP-2 (S1 Fig). [72]

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Fig 1. LOTR-1 contains both LOTUS and Tudor domains, and colocalizes with Z granules.

A) Top schematic depicts the location of the eLOTUS, mLOTUS and Tudor domain of LOTR-1. Bottom depicts the CRISPR edit used to add an N terminal 3X FLAG tag and GFP. Alleles generated for this study are indicated. B) Predicted 3D model overlap of the eLOTUS and mLOTUS domains of C. elegans LOTR-1 with the Oskar eLOTUS domain from Drosophila melanogaster. The α5 helix (highlighted) is only present in the eLOTUS domain. C) Predicted 3D structure of the Tudor domain (aa 534–670) of LOTR-1 overlapped with TD3 domain of mouse TDRD1. D) GFP::LOTR-1 and PGL-1::RFP in the germline of living worms. E) Super-resolution confocal imaging of GFP::LOTR-1 with PGL-1::RFP (top) and GFP::LOTR-1 with RFP::ZNFX-1 (bottom) in pachytene germ cells. F) Comparison of the center of mass difference from LOTR-1 for PGL-1 or ZNFX-1.

https://doi.org/10.1371/journal.pgen.1010245.g001

Default BLAST parameters fail to uncover the Tudor domain within D1081.7, but it readily appears in domain-enhanced searches and was previously identified using multiple sequence alignment. [12] The Tudor domain (aa 534–670) is most similar to Tudor domain 3 (TD3) of mouse TDRD1, which is known to bind symmetrically dimethylated arginine (sDMA) marks through a hydrophobic pocket that is created by the arrangement of four aromatic residues (Fig 1C, F767, Y774, F791, and Y794). [74] This pocket is stabilized by charge interactions between R775 and D793. The positions of these two residues are conserved in D1081.7 (R596 and D615). However, two of the four aromatic residues of the aromatic cage are replaced with other hydrophobic alternatives in D1081.7 (L613 and V616). How this impacts sDMA binding is unknown.

The combination of both LOTUS and Tudor domains in D1081.7 is unique to homologs of Tejas/TDRD5 and Tapas/TDRD7 in Drosophila and mammals, which are piRNA pathway-regulating components that localize to the perinuclear nuage of germ cells. D1081.7 is the only protein in C. elegans known to have both LOTUS and Tudor domains. The Tudor and LOTUS domains of TDRD7 and TDRD5 are arranged similarly to those of D1081.7, with TDRD5 sharing 34% sequence similarity to D1081.7 across 466 aa, and TDRD7 sharing 39% sequence similarity across 550 aa. While reciprocal BLASTP searches suggest that D1081.7 homologs are confined to class V nematodes (S1A), its domain architecture and predicted structure suggests that D1081.7 is a putative homolog of TDRD5 and TDRD7 in C. elegans (S1D). Given these structural features, D1081.7 was named LOTUS and Tudor domain protein 1 (LOTR-1).

To determine the expression of LOTR-1, CRISPR/Cas9 genome editing was used to place an N-terminal GFP tag on endogenous lotr-1 in a strain carrying PGL-1::RFP. LOTR-1 localized to germ granules throughout the adult germline; however, LOTR-1 granules appeared docked next to P granules marked by PGL-1::RFP (Fig 1D). Super-resolution confocal imaging of pachytene germ cells confirmed that LOTR-1 and PGL-1 granules appear adjacent to one another in the adult germline (Fig 1E, top), with their center of mass separated by an average of 284nm (Fig 1F). This pattern is similar to the P-granule docking of Z granules, Mutator foci, and SIMR foci, suggesting that LOTR-1 could reside within these germ granule sub-compartments. [70,71,75] To test this, LOTR-1 was examined for colocalization with RFP::ZNFX-1, which defines Z granules. Red and green channels show significant overlap (Fig 1E, bottom), with their center of mass separated by an average of only 71nm (Fig 1F). These results suggest that LOTR-1 partitions with Z granules instead of P granules in the adult germline.

Functional analysis of LOTR-1 domains reveals effects on subcellular localization and fertility

To understand how LOTR-1 is recruited to germ granules, several point mutations and deletions were introduced into the 3xFLAG::GFP::LOTR-1 strain (Fig 1A). Deletions of the mLOTUS and combined eLOTUS/mLOTUS domains do little to disperse truncated LOTR-1 from germ granules in young adults (Fig 2A) or reduce LOTR-1 granule number and size (S2A Fig). In contrast, the Tudor domain deletion disperses LOTR-1, decreasing LOTR-1 granule size and reducing the number of LOTR-1 granules in the adult germline by 85% (Figs 2A and S2A). A point mutation in the conserved arginine (R596C) of LOTR-1, which is predicted to stabilize the pocket with the potential to bind sDMAs, disperses LOTR-1 as well (Figs 2A and S2A).[37] These results suggest that the germ-granule association of LOTR-1 depends primarily on its Tudor domain, potentially mediated through well-characterized Tudor/sDMA interactions. In the absence of the Tudor domain, 37% of LOTR-1 granules are still retained during spermatogenesis in the fourth larval stage (Fig 2B, orange box, S2B Fig), suggesting that its retention in spermatogenic germ granules is not fully dependent on Tudor/sDMA interactions. One possibility is that when the nuclear envelope breaks down during spermatogenesis, causing germ granules to condense into fewer and larger aggregates, these larger granules facilitate LOTR-1 association even without its Tudor domain. Interestingly, lotr-1 mutants show only a subtle reduction in brood size in comparison to wild type, even after a generation grown at 26° C, in contrast to the robust temperature-sensitive sterile phenotypes of other P-granule mutants, suggesting a more specialized role for LOTR-1 in neighboring Z granules (S2C Fig).

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Fig 2. Germ-granule phenotypes in lotr-1 germlines.

A) Live-imaging of young adults shows the distribution of LOTR-1::GFP in the presence and absence of its LOTUS and Tudor domains. B) LOTR-1 distribution during spermatogenesis in the fourth larval stage (L4). Orange boxes indicate region of spermatogenesis. Live imaging of mCherry::PRG-1 and GFP::LOTR-1 in C) L4 stage and D) young adult germlines. E) Live imaging of RFP::ZNFX-1 in the germline of wild-type and lotr-1 mutant adults. F) Comparison of GFP::LOTR-1 and mCherry::PRG-1 expression in the germlines in znfx-1 mutant (top) and in lotr-1; znfx-1 double mutant (bottom) adults. G) RNAi depletion of mip-1 and mip-2 in GFP::LOTR-1 compared to empty vector control RNAi. H) lotr-1 RNAi compared to empty vector control RNAi in adult germlines of GFP-tagged LOTR-1, GLH-1, PGL-3, DEPS-1, MIP-1 and MIP-2 worms. Scale is 20 microns. Quantification provided in S2 Fig.

https://doi.org/10.1371/journal.pgen.1010245.g002

Tudor domains interact with PIWI Argonaute proteins via sDMAs. [76] To see if this interaction extends to C. elegans Argonautes, the distribution of the PIWI homolog PRG-1 was examined in the presence and absence of LOTR-1, and in a strain carrying LOTR-1 with the Tudor domain deletion. Under wild-type conditions, PRG-1 is constitutively associated with germ granules from the distal germline through gametogenesis, with more abundant expression during spermatogenesis. In both the lotr-1 full- and Tudor deletion backgrounds, 60–72% of PRG-1 granules persist during spermatogenesis, respectively (Fig 2C, orange box, S2B Fig). Only 29–42% of PRG-1 granules persist in pachytene germ cells, and 6%-28% in oocytes, in indicated lotr-1 mutant adults (Figs 2D and S2D and S2E). Both LOTR-1 and PRG-1 are deposited in residual bodies during spermatogenesis and absent from spermatids in the presence or absence of the Tudor domain of LOTR-1 (S2F Fig). These findings suggest that LOTR-1’s Tudor domain is not necessary to localize PRG-1 to germ granules, but the Tudor domain enhances this localization, potentially through interactions with sDMAs, more in oocytes than during spermatogenesis.

The impact of LOTR-1 is less pronounced on ZNFX-1 than it is on PRG-1 localization. In lotr-1 (deletion) germlines, 83% of ZNFX-1 granules persist in pachytene cells, and 61% persist in oocytes (Figs 2E and S2G and S2H). There was no significant difference in the number of ZNFX-1 granules in the lotr-1 LOTUS and Tudor deletion backgrounds, suggesting that the presence of LOTR-1 is not required for ZNFX-1 localization. An early frameshift mutation was introduced into ZNFX-1 to determine whether loss of ZNFX-1 reciprocally impacts GFP::LOTR-1 and RFP::PRG-1 expression. LOTR-1 localization remains largely intact in the znfx-1 mutant (Fig 2F). While PRG-1 localization looks wild-type in the first generation of znfx-1 homozygotes, only 66% (pachytene) and 54% (oocyte) of PRG-1 granules persist in subsequent generations (Figs 2F and S2D, S2E and S2 I). This is reduced to 32% (pachytene) and 16% (oocyte) in the lotr-1; znfx-1 double mutant, similar to levels in observed in lotr-1 single mutant, suggesting that the ability of lotr-1 and znfx-1 to disperse PRG-1 is not additive.

Next, LOTR-1 localization was examined in the absence of two other recently characterized LOTUS-containing proteins, MIP-1 and MIP-2. [72] MIP-1 and MIP-2 are MEG-3 interacting proteins that localize to P granules throughout development and promote germ granule condensation. Although MIP-1 and MIP-2 lack Tudor domains, they each contain two eLOTUS domains that could indicate some functional synergy with LOTR-1 (S1D Fig). RNAi depletion of mip-1 and mip-2 together has been shown to cause the dispersal of PGL-1, PGL-3, and GLH-1. [72] Similarly, GFP-tagged LOTR-1 granule numbers are reduced 41% and are less prominent around the nuclear periphery following mip-1; mip-2 RNAi, even though LOTR-1 is associated with Z and not P granules at this stage (Figs 2G and S2J). These results indicate that the LOTUS-domain proteins MIP-1 and MIP-2 directly or indirectly affect the localization of LOTR-1 at germ granules. GFP::LOTR-1 levels do not decrease with mip-1; mip-2 RNAi, suggesting that LOTR-1 granule loss is attributed to relocation and not depletion (S3H Fig). In contrast, MIP-1 and MIP-2 are not significantly dispersed from pachytene germ granules in lotr-1 mutants (S2J Fig). We sought to address whether LOTR-1 is required for localization of other germ-granule factors. GLH-1, PGL-3, DEPS-1 and MIP-1 GFP granules showed subtle, but non-significant, dispersal in the adult germline with lotr-1 RNAi, while MIP-2 granules were dispersed by 60% with lotr-1 RNAi, despite a slight increase in average MIP-2::GFP intensity (Figs 2H and S2K, S2L and S3H). This, together with the reciprocal effect of MIPs on LOTR-1 localization, suggests potential synergy between these three LOTUS-domain containing proteins.

LOTR-1 binding partners include ZNFX-1, germ-granule proteins, and the cytoskeleton

To explore the potential for direct associations between LOTR-1, GLH-1, MIP-1, MIP-2, and ZNFX-1 we performed yeast-two-hybrid (Y2H) analyses. MIP-1 and MIP-2 have previously been shown to both homo- and heterodimerize, as well as associate with GLH-1/Vasa through their N-terminal LOTUS domains. [72] Y2H confirmed associations between LOTR-1 and both MIP-1 and MIP-2 (S3A Fig), whereas no interaction between LOTR-1 and GLH-1 was detected (S3B Fig). Surprisingly, the association of MIP-1 and MIP-2 with LOTR-1 appears to be mediated through the C-terminal ends of these proteins and not through their LOTUS domains (S3A Fig). The Y2H also suggests that both MIP-1 and MIP-2 interact more strongly with LOTR-1 when one or both of its two LOTUS domains are removed (S3A Fig). Although Y2H does not show an interaction between LOTR-1 and the GLH-1 helicase, binding was confirmed between LOTR-1 and the ZNFX-1 helicase (S3C Fig). This suggests that distinct helicase/LOTUS-domain-protein pairs (GLH-1:MIP-1/2 and ZNFX-1:LOTR-1) exist in neighboring P and Z granules.

To complement the Y2H analysis, we took an unbiased, biochemical approach to find additional LOTR-1 interactions. The N-terminal 3xFLAG tag introduced into the endogenous LOTR-1 locus was used to immunoprecipitate LOTR-1 from both young adults and embryos for quantitative mass spectrometry (IP-qMS) (S3D Fig). Two independent anti-FLAG IP-qMS experiments were performed, both in young adults and embryos, which allowed us to identify proteins differentially enriched in 3xFLAG::GFP::LOTR-1 over 3xFLAG::GFP::lotr-1(deletion) IPs for both young adults and embryos (Fig 3A and 3B and S1 Table). Eight proteins, including LOTR-1, were enriched in all four LOTR-1 IP datasets, two embryo datasets and two young adult datasets (Fig 3A–3C). After LOTR-1, the most highly enriched among these was the ZNFX-1 helicase. The other six proteins are the germ granule-associated Argonautes PRG-1 and WAGO-1, the 3’UTR cleavage and stimulation factors SUF-1 and CPF-1, the histone deacetylase SIR-2.2, and F46G10.1. These eight proteins represent a central core of possible LOTR-1 interactions, reinforcing the role of LOTR-1 in germ granules and small RNA biogenesis.

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Fig 3. Quantitative IP-mass spectrometry analysis of LOTR-1 and ZNFX-1 immunoprecipitations.

A) Volcano plots show the significance and enrichment of proteins that immunoprecipitated with 3xFLAG::GFP::LOTR-1 over the lotr-1 deletion expressing GFP::3xFLAG alone, as identified by qMS. B) Venn diagram shows significantly enriched proteins that overlapped between two rounds of LOTR-1 IP-qMS from both embryo and young adult lysates. C) Table showing the change in LOTR-1 association in embryos and young adults when LOTUS or Tudor domains of LOTR-1 were deleted. D) Volcano plots show the significance and enrichment of proteins immunoprecipitated with 3xFLAG::GFP::ZNFX-1 over untagged ZNFX-1, as identified by qMS. E) Venn diagram shows significantly enriched proteins that overlapped between LOTR-1 and ZNFX-1 IP-qMS from both embryo and young adult lysates. F) Table showing the change in ZNFX-1 association in embryos and young adults when lotr-1 was mutated. For C and F, a purple gradient was applied to visualize fold increases (>1, darker) and decreases (<1, lighter) from the C) LOTR-1 and F) ZNFX-1 associations identified in the wild type (WT) replicates.

https://doi.org/10.1371/journal.pgen.1010245.g003

Expanding the interaction list by an additional ten proteins that showed significant enrichment in at least three out of the four embryo and young adult datasets, now includes the germ-granule proteins DEPS-1, the WAGO-4 Argonaute, and the YHTDC2-like DExH-helicase (F52B5.3) that was previously shown to interact with the germ-granule Argonaute CSR-1 (Fig 3B and 3C). [77] Of these 18 LOTR-1-interacting proteins, 5 are known to bind actin and regulate the cytoskeleton, including CAP-1, HUM-1, HUM-5, CLIK-1, and GSNL-1. [78–81] 12 of the 18 proteins originate from transcripts abundantly expressed in the germline (Fig 3C, bold), while transcripts encoding the remaining six (HUM-5, GSNL-1, F40A3.6, SIR-2.2, F46G10.1, and LGC-27) are lowly expressed in the germline and may reflect LOTR-1 interactions in whole-worm or embryo lysates that are not replicated in vivo. [82,83]

To distinguish if LOTR-1 associations obtained by co-immunoprecipitation (co-IP) are facilitated through its LOTUS or Tudor domains, LOTR-1 carrying deletions in either the e+mLOTUS (sam113) or Tudor (sam102) domains were immunoprecipitated from embryos and young adults and then analyzed with qMS to look for increased (>1, Fig 3C) or decreased (<1, Fig 3C) interactions. Anti-FLAG westerns confirmed expression of tagged LOTR-1 constructs at wild-type levels or above (S3E Fig). Deleting the LOTUS domains had a more pronounced effect than deleting the Tudor domain on the association of LOTR-1 with cytoskeletal proteins (reduced 80–100% in embryos, 50–70% in adults, Fig 3C, orange), 3’ UTR-cleavage and stimulation factors (reduced 60–70% in embryos, 30–50% in adults, Fig 3C, blue). LOTR-1 associations with other germ-granule proteins were decreased with both LOTUS and Tudor deletions (embryos and adults, Fig 3C, green). The effect was slightly more pronounced in the absence of the Tudor domain, which disperses LOTR-1 from germ granules. RFP::ZNFX-1 intensity levels are not significantly changed in lotr-1 mutants (S3H Fig). This suggests that the weakened interaction with LOTR-1 proteins bearing LOTUS or Tudor domain deletions is due to the absence of these domains, and not to altered ZNFX-1 protein levels in lotr-1 mutants.

Of the LOTR-1 associations specific to both embryo but not adult IPs, MEG-3 and WAGO-3 stand out because of their previously described association with germ granules. The association of LOTR-1 with MEG-3 in embryos is reduced 90% in the Tudor deletion, which shows this interaction is dependent on the Tudor domain of LOTR-1 (Fig 3C and S1 Table). Interestingly, the interaction with MEG-3 is enhanced almost two-fold in the absence of the LOTUS domains of LOTR-1. This may suggest that Tudor-dependent interactions between MEG-3 and LOTR-1 during embryogenesis are kept in check by other associations mediated through the LOTUS domains of LOTR-1. In turn, some LOTR-1 associations are more prominent in young adults. For example, MIP-2 associates with LOTR-1 in the young adult germline and this interaction primarily relies on the Tudor domain (Fig 3C and S1 Table). This is consistent with our Y2H results showing that MIP-2 interacts with the Tudor-domain-containing C-terminus of LOTR-1 (S3A Fig). Another of the more pronounced LOTR-1 interactors specific to young adults is the polo-like kinase PLK-3, and this interaction is disrupted when either the LOTUS or Tudor domains of LOTR-1 are removed (Fig 3C). The significance of these embryo and young adult-specific interactions warrants additional attention.

To confirm whether ZNFX-1 interactions reflect the core set of proteins bound to LOTR-1, anti-FLAG IP-qMS was performed on embryos and young adults expressing 3xFLAG::GFP::ZNFX-1 or an untagged control (Figs 3D and S3G). In addition to ZNFX-1, proteins enriched over untagged control in both embryo and young adult included LOTR-1 and its interactors (i.e., PRG-1, DEPS-1, WAGO-1, WAGO-4 and RME-2) (Fig 3D–3F). SMG-2, WAGO-3, and the 3’ UTR cleavage and polyadenylation factors SUF-1 and CPF-1 were enriched in embryos only, while MIP-2 and PLK-3 were again only enriched in young adults (Fig 3E and 3F). These results support a substantial overlap of interactions between ZNFX-1 and LOTR-1 within Z granules. To determine which of these ZNFX-1 associations depend on LOTR-1, ZNFX-1 IP-qMS was performed in both embryo and adult lotr-1(xf58) mutants, which harbor an in-frame deletion that removes most of the LOTUS domain (Figs 3F and 1A). The association of ZNFX-1 with PRG-1, DEPS-1, WAGO-1, WAGO-3 and WAGO-4 is reduced 40–70% in the lotr-1 mutant embryos, while its association with SUF-1, CPF-1, and SMG-2 in embryos is practically eliminated (Fig 3E and 3F and S2 Table). This suggests that LOTR-1 stabilizes ZNFX-1 interactions with Argonaute proteins while acting as the link between ZNFX-1 and mRNA/3’UTR binding factors. Interestingly, the LOTUS-containing protein MIP-2 increases its association with ZNFX-1 two-fold in lotr-1 mutant adults, which may support a compensatory function for MIP-2 and LOTR-1.

lotr-1 mutants deregulate subsets of 26G- and 22G-RNAs and display changes in 22G-RNA coverage

Having established that LOTR-1 interacts with ZNFX-1 in Z granules, we sought to compare mutant phenotypes. Mutations affecting small RNA biogenesis and amplification frequently exhibit transgenerational sterility or mortal germline (Mrt) phenotypes. [40,84] To address if LOTR-1 is required for transgenerational germline health, fertile generations were counted until sterility ensued for three different lotr-1 mutant alleles (S4A Fig). Each of three lotr-1 alleles failed to propagate beyond 50 generations, while wild-type (N2) worms remained fertile. Because znfx-1 mutants have transgenerational epigenetic inheritance defects that manifest after several generations at 25° C [70], brood sizes of single and double lotr-1; znfx-1 mutants were compared to wild-type broods over the course of five generations (S4B Fig). While brood sizes are similarly and significantly decreased in both lotr-1 and znfx-1 mutants by the fifth generation, the effect was not additive, again suggesting that LOTR-1 and ZNFX-1 act in the same pathway.

The Mrt phenotype and impact on germ granules observed in lotr-1 suggest these mutants are defective in some aspect of small RNA biogenesis. Since many LOTUS domain proteins have functions in piRNA biology, [76] we asked if this is the case for LOTR-1 using lines that carry a 21U-RNA/piRNA sensor construct. This transgene contains a reporter for GFP::H2B expression that has been silenced through a piRNA target site in its 3’UTR; mutations affecting piRNA biogenesis or secondary 22G-RNA production de-silence expression of this transgene. [45] Loss-of-function alleles of lotr-1 were crossed into two different sensor lines carrying this transgene: one in which silencing is still dependent on 21U-RNAs (S4C Fig), and another under stable RNAe that is maintained independent of the initial 21U-RNA trigger (S4D Fig). Mutations in lotr-1 were unable to activate either piRNA sensor strain (S4C and S4D Fig), showing that LOTR-1 is not required for 21U-RNA-dependent silencing or for RNAe.

As described in the introduction, C. elegans expresses a variety of small RNA species. [40] To address if LOTR-1 is required for some aspect of small RNA biogenesis, we sequenced small RNAs in wild-type and in lotr-1(sam117) (with the entire lotr-1 coding sequence deleted, see Fig 1A). Small RNA sequencing in gravid adults showed that levels of 21U-RNAs and miRNAs are not significantly changed in lotr-1 mutants (Fig 4A and 4B). The 26G-RNA pool is significantly affected and fifty genes show depletion of 26G-RNAs (Fig 4A and 4B). Of these, all but two are annotated to be targets of the known effectors ERGO-1, NRDE-3, or ALG-3/4, but the majority are ERGO-1 targets (47/50, S3 Table). [63,66,85] A GFP::NRDE-3 transgene, which localizes from the nucleus to the cytoplasm upon disruption of the 26G-RNA pathway, [63,86] remained expressed in the nucleus when crossed into lotr-1 mutants (S4F Fig), indicating the 26G-RNA pathway is not critically impaired. It has been recently reported that deregulation of mutator-dependent 22G-RNAs targeting the eri-6/7 locus may destabilize ERI-6/7, a protein required for ERGO-1-dependent 22G-RNA biogenesis [87]. In lotr-1 mutants, we find a mild increase of 22G-RNAs in this locus (S4G Fig), which may explain the lower levels of ERGO-1-dependent 26G-RNAs (Fig 4A). Taken together, these results suggest that while LOTR-1 is not absolutely required for normal 26G-RNA biogenesis and silencing, a subset of 26G-RNA targets is affected by the depletion of LOTR-1.

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Fig 4. 26G- and 22G-RNA deregulation in lotr-1 mutants.

A) Relative levels of the indicated small RNA populations normalized to all non-structural reads. B) Differential expression analysis to determine genes and transposons that are significantly depleted or enriched of mapped small RNAs in lotr-1 mutants. The MA plots depict DESeq2 differential results for miRNAs (n = 257), 21U-RNAs over 21ur loci (n = 14328), and 22G-/26G-RNAs over protein coding genes (n = 20222), lincRNAs (n = 172), pseudogenes (n = 1791) and transposons (n = 151); significant changes (>2-fold at 10% FDR) are colored in red with the number of up- and down-regulated hits indicated. C-E) Metagene plots depicting relative (normalized per individual gene as in [71], upper panels) as well as total (lower panels) 22G-RNA coverage over genes whose 22G-RNAs are up-regulated, down-regulated or unaffected in the lotr-1 mutants. Note that the total (cumulative) coverage plots utilize DESeq2-based normalization for library composition and are biased towards genes with strong 22G-RNA levels. TSS/TES, transcription start/end site.

https://doi.org/10.1371/journal.pgen.1010245.g004

Global 22G-RNA levels are not changed overall; however, hundreds of genes have deregulated 22G-RNA levels in lotr-1 mutants (Fig 4A and 4B and S3 Table). Next, we asked if the deregulated 22G-RNAs map to particular gene classes. In lotr-1 mutants, the majority of genes with deregulated 22G-RNAs are protein-coding genes (S5A Fig). 67 transposons display significantly up-regulated 22G-RNAs (S5A Fig and S3 Table), e.g. aggregated data for Tc1 transposons show approximately 3-fold increased 22G-RNA levels in lotr-1 mutants (FDR-adjusted p-value of 2.13E-95, S3 Table). However, a reporter for Tc1 transposon mobility was not affected by the lotr-1 mutation, suggesting that the upregulation of Tc1 22G-RNAs observed in lotr-1 animals does not disrupt Tc1 silencing (S4E Fig). After separating 22G-RNAs into subgroups according to their Argonaute cofactors and target genes [40], we find that most genes with deregulated 22G-RNA levels in lotr-1 mutants belong to the mutator and/or WAGO classes (S5B Fig and S3 Table). Overall, these results suggest that hundreds of genes show deregulated 22G-RNA levels, both increased and decreased, in lotr-1 mutant animals across a range of gene classes and 22G-RNA targets.

In znfx-1 mutants, 22G-RNA coverage is altered, revealing a shift toward the 5’ end of the target RNAs. [71] Metagene analysis of 22G-RNA coverage along transcripts showed a similar effect in the genes with deregulated 22G-RNA levels in lotr-1 mutants, with stronger 5’ shift in the up-regulated genes (Figs 4C–4E and S5C and S5D). WAGO and mutator targets showed a similar 5’ shift of 22G-RNAs in lotr-1 mutants compared to wild-type (S5C–S5F Fig). This 5’ shift of 22G-RNAs is not observed across all loci with robust 22G-RNA targeting (S5G Fig). Unlike znfx-1, the 5’ shift in 22G-RNA coverage in lotr-1 mutants is observed in WAGO/mutator, but not in CSR-1 targets (S5H Fig). Notably, CSR-1 was not enriched in LOTR-1 IP-qMS, in either the embryo or young adult lysates. 22G-RNA coverage of ALG-3/4 targets is affected mainly in the 5’ region, whereas ERGO-1 targets are not affected in lotr-1 mutants (S5I–S5J Fig). Of note, similar 22G-RNA deregulation and changes in 22G-RNA distribution across WAGO/mutator targets are also observed upon sequencing small RNAs from other lotr-1 mutant alleles with partial deletions in the coding sequence (S5K–S5M Fig). Therefore, we conclude that LOTR-1 functions with ZNFX-1 to balance the production of 22G-RNAs across WAGO/mutator, but not CSR-1, target RNAs.

LOTR-1 affects the inheritance of RNAi

The deregulation of 22G-RNA coverage over WAGO/mutator targets observed in lotr-1 mutants, and the interaction of LOTR-1 with ZNFX-1 and WAGO proteins, may point to defects in exogenous RNAi and its inheritance. To test whether this is the case, the incidence of embryonic lethality following pos-1 RNAi was examined in wild type and two lotr-1 alleles, using a mut-7 mutant as a control known to be resistant against pos-1 RNAi (Fig 5A). Embryonic lethality was mildly suppressed in the full deletion allele, but not to the level observed in mut-7 (Fig 5A, left). Next, we tested for defects in the inheritance of RNAi, using RNAi against GFP in wild type and lotr-1 mutants carrying an H2B::GFP reporter. GFP RNAi was 100% effective in knocking down GFP expression in both wild-type and lotr-1 worms. In contrast to znfx-1 mutants [70], lotr-1 mutants faithfully inherited the GFP silencing to the F1 generation (Fig 5B). In a separate experiment, inheritance of H2B::GFP silencing in lotr-1 deletion mutants was followed for ten generations in isolated lineages, which revealed that lotr-1 mutants can display enhanced RNAi inheritance (Fig 5C). Thus, although LOTR-1 and ZNFX-1 interact and the respective mutants share phenotypes, they clearly have distinct functions.

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Fig 5. RNAi sensitivity and inheritance in lotr-1 mutants.

A) Sensitivity of two different lotr-1 mutant alleles against pos-1 RNAi, using mut-7(xf125) mutants as a known pos-1 RNAi resistant control. Hatching proportion (y-axis) was scored; horizontal blue lines represent the fitted hatching probabilities. P-values were determined using Likelihood Ratio Tests. Error bars: bootstrap 95% confidence intervals for hatching probabilities. B) Inheritance of RNAi was tested using RNAi against GFP, targeting a transgene expressing a GFP::histone2B fusion (mjIS31) in oocytes [50]. Silencing in the treated P0 generation was fully effective, as was the inheritance of silencing to the F1. C) Transgenerational inheritance assay using RNAi against a GFP::histone2B transgene (ruIs32) in WT and the deletion allele lotr-1(usa1) revealed enhanced silencing up to 10 generations in the absence of LOTR-1. D) A model summarizing how LOTR-1, together with ZNFX-1, could affect 22G RNA production from an RdRP-targeted transcript. We note that all interactions are based on IP-qMS and Y2H experiments. Since these techniques do not report on whether a given interaction reflects a stable complex, or rather a more transient type of interaction, further study will be required to define the precise composition and nature of the associated molecular machinery.

https://doi.org/10.1371/journal.pgen.1010245.g005

Sequence, structural, and evolutionary analysis

To characterize LOTR-1 and its interactions, we used sequence analysis (PSI/Delta-BLAST, HHPred, and multiple sequence alignment with PRALINE) to identify conserved protein domains. [95–98] Homology modeling was performed with RaptorX: eLOTUS domain was constructed from Oskar LOTUS (PDB ID: 5NT7); mLOTUS domain from Cdt1 (PDB ID: 5MEC), which contains winged-helix domains that interact with subunits of the MCM helicase motor; and Tudor domain from TD3 (PDB ID: 4b9wA) from TDRD1.

Conservation of LOTR-1 in nematodes was analyzed as previously described. [99] Proteomes of nematode species were downloaded from WormBase ParaSite (WBPS16) and BLAST databases were created from each proteome with makeblastdb–dbtype prot. Then, two sets of whole-proteome BLASTP searches were performed: 1) using the C. elegans proteome as query versus all other proteomes; and 2) the reciprocal BLASTPs using all the proteomes as query versus the C. elegans proteome. From the first set of BLASTP searches, we extracted the best BLAST hit for C. elegans D1081.7/LOTR-1 (full length) in other species. We used the best BLAST hits, to get a bit score from the reciprocal BLASTP (second set of BLASTPs) when D1081.7/LOTR-1 was the best reciprocal BLASTP hit. The bit scores were normalized by dividing by the C. elegans bit score. When no reciprocal BLAST hit was found, the score of 0 was given. The scores were loaded into R v.4.1.0 and plotted using publicly available packages (ggplot v.3.3.5, using geom_tile()). Nematode clades were annotated similar to Rodrigues et al., 2019. [99]

Strain generation and maintenance

C. elegans strains were maintained using standard protocols. [100] A complete list of strains and alleles generated for this study is provided (S4 Table). CRISPR/Cas9 was used to place tags on endogenous genes and generate mutant alleles as described. [101] All alleles generated for this study were sequence verified.

Imaging

Live worms were mounted on agarose pads in egg buffer (25 mM HEPES, 120 mM NaCl, 2 mM MgCl2, 2 mM CaCl2, 50 mM KCl, and 2 mM levamisole), covered by No. 1.5 coverslips, and imaged with a Leica DMI6000B widefield epifluorescence microscope equipped with a Leica HC PL APO 63x/1.47NA oil immersion objective. Samples were excited with a Leica EL6000 light source and EGFP (11525314) and Texas Red filter (11525310) sets. Images were acquired with a Leica DFC9000GT camera controlled with LAS acquisition software with the same conditions, 2048X2048 pixel and in 16 bits. Figs 1D, 2A–2F and S2F and S2I are deconvolved 30μm projections of the germline loop region. For quantification in S2A,B,D,E,G and H, a single-plane was acquired. Fig 1E images are 8μm and 1μm maximum intensity projections of pachytene germ cells acquired with an upright Zeiss LSM 980 confocal microscope (axio examiner) equipped with AiryScan2 detector and a Zeiss Plan Apochromat 63X/1.4NA oil immersion objective, mounted on a piezo insert and controlled by Zen Blue 3.1 acquisition software. RFP and EGFP fluorescence were respectively excited with the 561nm (2.5% laser power) and 488 nm (1.50% laser power) laser line and the emission were collected from 573 to 627nm and from 499 to 557nm. A 8um Z-stack series optical sections were collected sequentially, in bidirectional mode, with a pixel size of 0.043um X 0.043um, a step-size of 0.17 um, at zoom 12, in 16 bit and in Super Resolution mode (SR). Images were processed with a strength of the deconvolution set to 8.5 or 6.7 for EGFP and 8.1 and 6.3 for RFP 8.1 and saved in CZI file[102]. Image analysis was performed using the open-source FIJI software. [102]

Center of mass differences were calculated from 20x40 micron red/green sections from single-plane images of pachytene cells, 1 image/worm, 20 worms/strain. A FIJI macro (included in S5 Table) was applied that uses a Difference of Gaussian (DoG) algorithm to identify and mask germ granules in each channel (https://micro.magnet.fsu.edu/primer/java/digitalimaging/processing/diffgaussians/index.html). The JACoP plugin was used to identify the geometrical center for each germ granule and then calculate the ‘distance-based colocalization’ with its nearest neighbor in the corresponding channel. [103] S5 Table contains the distances between granule pairs in each of the twenty images from both strains that were examined.

Germ-granule size and number were calculated from 20x40 micron sections from single-plane images of pachytene, oocyte, or spermatogenesis regions, 1 image/worm, 20 worms/strain. To do this, an ImageJ ‘germ-granule counter’ macro (include in S5 Table) was applied to ‘MaxEntropy AutoThreshold’ granules and ‘Analyze Particles’, providing granule number and size in each 20x40 section. S5 Table contains granule number and size data for the graphs in S2 Fig. RFP::ZNFX-1, DEPS-1::GFP, MIP-1::GFP, and MIP-2::GFP levels in S3H Fig were calculated from the average pixel intensity of 20x40 micron sections from single-plane images of the pachytene region, 1 image/worm, 20 worms/strain.

RNAi

RNAi feeding was performed as previously described. [104] The L4440 plasmid in HT115 bacteria was used as the RNAi control. RNAi was performed starting on L1 worms, with at least three biological replicates, and used feeding constructs previously described. [105] For simultaneous depletion of mip-1 and mip-2, equal parts from the Ahringer RNAi feeding library clones were mixed and fed to L1 worms.

pos-1 RNAi resistance assays: 10 L3 stage animals were singled onto pos-1 RNAi feeding plates. After two days, adults were removed and eggs were counted, and the number of hatched larvae counted the following day. Data were analyzed with a generalized linear mixed effects model for Bernoulli-distributed binary data using the R package lme4. [106] The log-odds-ratio of hatched larvae was modeled as a linear combination of coefficients for three factors: the focal factors genotype and RNAi treatment were modeled as fixed effects, while plate membership was included as a random effect to account for the (in some cases) considerable differences in hatching frequencies between plates of the same genotype-condition group. Likelihood ratio tests were used to determine significance of differences in RNAi treatment effect between WT and lotr-1(xf60) and between WT and lotr-1(sam117). 95% confidence intervals for the hatching probabilities of the respective genotype-treatment groups (excluding random plate effects) were calculated by parametric bootstrapping (bootstrap size 10000).

Transgenerational epigenetic inheritance (TEI) assays: Two sets of experiments were performed in different laboratories to test TEI.

For Fig 5B, Worms expressing GFP::H2B (mjIS31) in wild-type N2 and lotr-1(xf60) mutants were bleached and embryos were placed on E. coli HT1115 gfp RNAi or control L4440 plates. [50,86] After four days, adult RNAi treated P0 worms were screened for presence or absence of GFP expression. The RNAi treated P0 worms were then bleached and F1 embryos were placed onto OP50 plates. Four days later F1 worms were screened for GFP expression. We considered worms that expressed GFP at levels comparable to non-RNAi treated worms as GFP expressing, and worms that were either completely silenced or had barely detectable expression of GFP as silenced. To score GFP expression we used a Leica Thunder Imager Live Cell microscope with a HC FLUOTAR L VISIR 25x/0.95 water objective. Representative images were taken with a Leica AF7000 microscope with a sCMOS Flash 4.0 Hamamatsu camera and a HCX PL APO 40x/1.1 water objective using LASX Application Suite X 3.7.4.23463 software. Image J v. 1.53c was used for visualization. For microscopy sample preparation, approximately 50 worms were washed in a drop of M9 buffer and subsequently transferred to a 50 μl drop of 40mM sodium azide in M9 buffer on a coverslip. Once the worms were paralyzed, excess buffer was removed and a glass slide with a 2% (w/v) Agarose pad was placed on top of the coverslip.

For Fig 5C, L1 animals expressing H2B::GFP (ruIs32) in wild-type N2 and lotr-1(usa1) mutants were placed on GFP(RNAi) and control L4440(RNAi) plates at 20°C. 100 progeny from each plate were scored for GFP expression in the germ line using a Leica DM5500 widefield microscope with a 40X objective, and the Leica LASX software. From the GFP(RNAi) plates, ten silenced young adults from both wild type and lotr-1 mutants were singled to individual plates seeded with OP50 on NGM plates (referred to as “F1 generation” in Fig 5C). For each line, the progeny (n>50) was scored for GFP expression. Lines were recorded as still silenced if no GFP or only weak GFP expression was detected. Silenced lines were propagated by moving 3 young adults into a fresh NGM plate every new generation, and scored for GFP expression for a maximum of ten successive generations. Data from three replicate experiments were pooled and the proportion of silenced lines in each generation, out of a total of 30 individual lines, was plotted. Kaplan-Meier curves were fitted to assess survival and significance tested with a log-rank test.

Brood size

Brood sizes were counted from each strain for S2C Fig after placing ten or more L4 worms on individual plates at 20°C and 26°C. For the generational brood size assay in S4B Fig, 5 L4 worms from each strain were cloned to individual plates and shifted from 20°C to 25°C. Worms were kept at 25°C for 5 generations, with L4 progeny from each plate used to start the subsequent generation.

Germline mortality

Worms were assessed for the mortal germline phenotype using the assay described in Ahmed et al. 2000. [107] Three lotr-1 strains (with the xf58, xf60, xf61 alleles) and wild-type N2 animals were synchronized by bleaching and overnight hatching in M9 and L1-arrested animals. Five L1-L2 worms were picked to a new plate and grown at 25°C. The sixth day L1-L2 worms, corresponding to the third generation from the picked worms, were again hand-picked to a new plate. Fertile generations were counted until sterility ensued and no progeny could be isolated. Five replicates were used in this assay, N2 worms were used as control. Worms were always picked before starvation to avoid any effect it might cause during the assay, including extension of transgenerational germline lifespan. [108]

Yeast two hybrid

Full-length or truncated cDNAs of mip-1, mip-2, glh-1, lotr-1 and znfx-1 and DNA encoding 3xFLAG tag were cloned in-frame with GAL4-AD or GAL4-BD into the multiple cloning site of pGAD-C1 and pGBD-C1. [109] Yeast cells from strain PJ69-4a were transformed with 1 μg of plasmid and carrier DNA with lithium acetate [110].Transformed cells were then plated on the appropriate drop-out media, and the presence of the newly introduced plasmids was checked by colony PCR. Bait-prey interactions were done on SC-Leu-Trp-Ade plates or SC-Leu-Trp-His plates supplemented with varying concentrations of 3-amino-1,2,4-triazole, as previously described [72].

Reporters for piRNA activity and NRDE-3 localization

Experiments were performed as previously described. [45,63,86] Crosses (S4C-D) were set up in 6 cm Petri dishes with only 10 μl of OP50 to favor nematode meeting and mating. Wide-field fluorescence microscopy images (S4C-D, G) were acquired and processed with the Leica Application Suite (LAS) software (v3.1.0) on a Leica DM6000B.

Tc1 reversion

Tc1 reversion experiments were performed as previously described. [111] Crosses were set up, and cross-progeny genotyped as described for the sensor experiments above. Twitcher unc-22(st136) worms homozygous for lotr-1 were grown at 25°C: 24 independent populations of lotr-1; unc-22(st136) per lotr-1 allele. Controls and lotr-1 populations were grown in parallel for several generations, namely four plate passages, corresponding to 8–12 generations. Controls were grown in 2 independent populations for the same duration of time. Plates were screened every 2–3 days for revertants, i.e., animals that lost the twitching phenotype.

Small RNA sequencing and bioinformatic analysis

α-FLAG Immunoprecipitation and Western Blot

Immunoprecipitation and mass spectrometry were performed largely as described. [63] Animals were grown either in normal OP50-seeded NGM plates or on OP50 high-density plates. The protocol for the production of the latter was adapted from (Schweinsberg and Grant, 2013, https://www.ncbi.nlm.nih.gov/books/NBK179228/). In short, commercially available chicken eggs were cracked, the yolks isolated and thoroughly mixed with LB medium (50 mL per egg yolk). Then, the mix is incubated at 65°C for 2–3 hours. After cooling down, pre-grown OP50 liquid culture is added to the mix (10 mL per egg). 10 mL of this preparation is poured into each 9 cm NGM plate and plates are decanted the next day. After 2–3 days of further bacterial growth and drying, plates should be stored at 4°C.

Worms expressing 3xFLAG-tagged LOTR-1 and ZNFX-1 were grown and synchronized by bleaching and overnight hatching in M9. Synchronized young adults with no visible embryos were collected 51–56 hours post-plating, washed in M9, followed by a last wash in water, and frozen in dry ice. Prior to IP, samples were thawed, 2x Lysis buffer was added (50 mM Tris/HCl pH 7.5, 300 mM NaCl, 3 mM MgCl₂, 2 mM DTT, 0.2% Triton X-100, 1 complete Mini protease inhibitor tablet, #11836153001) and lysis was conducted with sonication in a Bioruptor Plus (Diagenode, on high level, 10 cycles of 30 seconds on/off). Embryos were collected by bleaching gravid animals approximately 72 hours after plating, washing with M9, plus one last washing step in 1x lysis buffer (25 mM Tris/HCl pH 7.5, 150 mM NaCl, 1.5 mM MgCl₂, 1 mM DTT, 0.1% Triton X-100, 1 complete Mini protease inhibitor tablet, #11836153001) and by freezing worm pellets in liquid N₂ in a pre-cooled mortar. Prior to IP, the pellets were ground to a fine powder in a pre-cooled mortar, then transferred to a cold glass douncer and sheared for 40 strokes with piston B. First round of IPs to FLAG-tagged LOTR-1 were performed using 1.5 mg of total embryo or young adult extracts, while for the second round of these experiments 1 mg of embryo or young adult extracts were used. IPs to FLAG-tagged ZNFX-1 were performed using 1 mg of young adult extract and 0.65 mg of embryo extract. IPs were performed in quadruplicates, with exception of FLAG IPs to LOTR-1(ΔTudor) in embryos, which were conducted in triplicate. FLAG-tag immunoprecipitation was performed using an α-FLAG antibody (Monoclonal ANTI-FLAG M2 antibody produced in mouse, Sigma Aldrich, #F3165) and Protein G magnetic beads (Invitrogen Dynabeads Protein G; #10004D). 30 μL of beads were used per IP. Beads were washed three times with 1 mL of Wash Buffer (25mM Tris/HCl pH 7.5, 300 mM NaCl, 1.5 mM MgCl₂, 1 mM DTT, 1 complete Mini protease inhibitor tablet, #11836153001). 2 μg of α-FLAG antibody were incubated with the beads and the extract for approximately 3 hours, rotating at 4°C. Afterwards, samples were washed 5 times with 1 mL of Wash Buffer. Finally, the beads were resuspended in 1x LDS buffer (Thermo Scientific, #NP0007) supplemented with 100 mM DTT and boiled at 95°C for 10 minutes.

Samples for mass spectrometry were prepared in quadruplicates: four lysates were prepared from pelleted young adult worms or pelleted embryos for each strain. Independent anti-FLAG IP-qMS experiments were then performed on each of these lysates. This same process was then duplicated on a separate occasion with new young adult and embryo lysates, again using quadruplicates.

Western blot in S3D Fig was performed exactly as described [113] using a mouse monoclonal anti-FLAG antibody (Sigma-Aldrich, #F3165) in a 1:5000 dilution in Skim Milk solution (1x PBS, 0.1% Tween-20, 5% (w/v) Skim Milk Powder), and a rabbit anti-Actin antibody (Sigma-Aldrich, #A2066) in a 1:500 dilution in Skim Milk Solution. We first probed the IP bait with a HRP-linked secondary antibody (Cell Signaling Technology, anti-mouse IgG, #7076; 1:10,000 dilution in Skim Milk Solution), and Actin was subsequently probed with a LI-COR secondary antibody (LI-COR IRDye 800CW Donkey anti-Rabbit IgG (H+L), #926–32213, in a 1:15,000 dilution in Skim Milk Solution). 1.5 mg of young adult extract were used for LOTR-1 anti-FLAG IPs, while 1 mg of young adult extract were used for ZNFX-1 anti-FLAG IPs.

The western blot in S3E Fig was performed as above using fed and synchronized young adults washed from two plates (approximately 1000 worms/sample), flash frozen in 50 μl of 2X Laemmli sample buffer (65.8mM Tris-HCl, pH 6.8, 2.1% SDS, 26.3% (w/v) glycerol, 0.01 bromophenol blue). Samples were run on a 4–15% gradient gel (BioRad), simultaneously probed with anti-FLAG antibody (1,5000) and a mouse anti-tubulin antibody (Sigma-Aldrich, #T9026 in a 1:3000 dilution). FLAG and tubulin were detected with HRP-linked secondary (goat anti-mouse antibody Invitrogen #31430 in a 1:3000 dilution). ImageJ was used to measure relative expression levels of LOTR-1 in relation to tubulin.

The Western blot in S3F Fig was performed using extracts derived from roughly 8000 synchronized young adult animals raised on the appropriate RNAi plates. Collected animals were cleared of the bacteria and flash frozen in RIPA buffer (150 mM NaCl, 1% v/v Triton X-100, 0.5% w/v sodium deoxycholate, 0.1% w/v sodium dodecyl sulfate, and 0.05 M Tris-HCl pH 7.5) supplemented with 1% v/v protease inhibitors (Sigma P8340), then homogenized with 425–600 μm diameter glass beads (Sigma G8772). Westerns were run with 80 μg of supernatants per lane on 4–12% bis-tris gels (Thermo Fisher Scientific NP0335) and simultaneously probed with anti-FLAG (Sigma clone M2 F3165, 1:4000) and anti-tubulin (DSHB clone 12G10, 1:10000), then detected with an HRP-linked secondary serum (GE NXA931, 1:10000). ImageJ was used to measure relative expression levels of LOTR-1 in relation to tubulin.

Mass spectrometry

IP samples were boiled at 70°C for 10 minutes and separated on a 4–12% gradient Bis-Tris gel (Thermo Fisher Scientific, #NP0321) in 1x MOPS (Thermo Fisher Scientific, #NP0001) at 180 V for 10 minutes. Then, samples were processed separately, first by in-gel digestion [114,115], followed by desalting with a C18 StageTip. [116] Afterwards, the digested peptides were separated on a heated 50-cm reverse-phase capillary (75 μm inner diameter) packed with Reprosil C18 material (Dr. Maisch GmbH). Peptides were eluted along a 90 min gradient from 6 to 40% Buffer B (see StageTip purification) with the EASY-nLC 1,200 system (Thermo Fisher Scientific). Measurement was done on an Orbitrap Exploris 480 mass spectrometer (Thermo Fisher Scientific) operated with a Top15 data-dependent MS/MS acquisition method per full scan. All raw files were processed with MaxQuant (version 1.5.2.8) and peptides were matched to the C. elegans Wormbase protein database (version WS269). Differential enrichment was defined for 3xFLAG::GFP::LOTR-1 IPs versus 3xFLAG::GFP::lotr-1(deletion) IPs. Venn diagrams were produced with Venny 2.1 (https://bioinfogp.cnb.csic.es/tools/venny/). Raw data and detailed MaxQuant settings can be retrieved from the parameter files uploaded to the ProteomeXchange Consortium via the PRIDE repository: identifier PXD025186.