IUPred2A (Mészáros et al., 2018) predicts in the MEG-3 sequence a N-terminal domain with high disorder and a C-terminal domain with lower disorder separated by a boundary region with mixed order/disorder (aa544–698) (Figure 1A, B). The C-terminus contains a predicted ordered 44 amino acid sequence (aa700–744) with homology to the HMG-like-fold found in the GCNA family of intrinsically disordered proteins (Figure 1C). Like MEG-3, GCNA family members contain long N-terminal disordered domains, but these do not share sequence homology with the MEG-3 IDR (Carmell et al., 2016). To test the functionality of MEG-3 domains in vivo, we used CRISPR genome editing to create four MEG-3 derivatives at the endogenous locus: MEG-3_Cterm (aa545–862); MEG-3_IDR (aa1–544); MEG-3₆₉₈, an extended version of MEG-3_IDR terminating right before the HMG-like motif; and MEG-3_HMGL-, a full-length MEG-3 variant with alanine substitutions in four conserved residues in the HMG-like motif (Figure 1B, C). (We also constructed a MEG-3_Cterm (aa545–862) variant with mutations in the HMG-like motif, but this variant was not expressed at sufficiently high levels for analysis.) The MEG-3 variants were created in a C. elegans line where the meg-4 locus was deleted to avoid possible complementation by MEG-4, a close MEG-3 paralog. To allow visualization of MEG-3 protein by immunofluorescence, each variant (and wild-type meg-3) was tagged with a C-terminal OLLAS peptide (Figure 2A). We avoided the use of fluorescent tags as fluorescent tags have been reported to affect the behavior of proteins in P granules (Uebel and Phillips, 2019).

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Condensation and enrichment of MEG-3 in four-cell embryos.

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As reported previously for untagged MEG-3 (Wang et al., 2014), MEG-3 tagged with OLLAS could be detected diffusively in the cytoplasm and in condensates (Figure 2A). Before polarization, MEG-3 was uniformly distributed throughout the zygote. After polarization, MEG-3 in the cytoplasm and in condensates became enriched in the posterior half of the zygote destined for the germline blastomere P₁ (‘germ plasm’). MEG-3 continued to segregate preferentially with P blastomeres in subsequent divisions (P₁ through P₄) (Figure 2A).

All four MEG-3 variants exhibited unique localization patterns distinct from wild-type. MEG-3_IDR enriched in posterior cytoplasm and segregated preferentially to P blastomeres but did not appear robustly in condensates until the four-cell stage (P₂ blastomere, Figure 2A). MEG-3₆₉₈ behaved identically to MEG-3_IDR (Figure 2—figure supplement 1A). MEG-3_Cterm did not enrich asymmetrically in the cytoplasm but formed condensates in the zygote posterior and continued to form condensates only in P blastomeres despite being present in the cytoplasm of all cells (Figure 2A). MEG-3_HMGL- behaved most similarly to wild-type MEG-3 enriching in the zygote posterior and forming condensates as early as the one-cell stage, although the condensates appeared smaller at all stages (Figure 2A).

For each MEG-3 derivative, we quantified the number of condensates and the degree of enrichment in the P blastomere (P₂) over somatic blastomeres and in condensates over the cytoplasm at the four-cell stage. Wild-type MEG-3 and MEG-3_HMGL- formed a similar number of condensates, while MEG-3_Cterm formed fewer and MEG-3_IDR the least in the four-cell stage (Figure 2B). The MEG-3_Cterm did not enrich in the P₂ blastomere, whereas MEG-3_IDR and MEG-3_HMGL- enriched as efficiently as wild-type (Figure 2C). Finally, none of MEG-3 derivatives enriched in condensates as efficiently as wild-type (Figure 2D).

After the four-cell stage, the low levels of wild-type MEG-3 and MEG-3_HMGL- inherited by somatic blastomeres were rapidly cleared. In contrast, MEG-3_IDR and MEG-3_Cterm persisted in somatic blastomeres at least until the 28-cell stage (Figure 2—figure supplement 1B). Western analyses revealed that MEG-3 and MEG-3_HMGL- accumulate to similar levels, whereas MEG-3_IDR and MEG-3_Cterm were more abundant in mixed-stage embryo lysates, consistent with slower turnover in somatic lineages (Figure 2—figure supplement 1C).

The condensation, segregation, and turnover patterns of MEG-3, MEG-3_IDR, MEG-3_Cterm,and MEG-3_HMGL- are summarized in Figure 2E. From this analysis, we conclude that (1) the MEG-3 IDR is necessary and sufficient for enrichment of cytoplasmic MEG-3 in germ plasm, (2) the MEG-3 C-terminus is necessary and sufficient to assemble MEG-3 condensates in germ plasm starting in the zygote stage, (3) the HMG-like motif enhances, but is not essential for, condensation, and (4) both the C-terminus and the IDR are required for timely turnover of MEG-3 in somatic lineages.

MEG-3 and MEG-4 are required redundantly to localize PGL condensates to the posterior of the zygote for preferential segregation to the P lineage (Smith et al., 2016; Wang et al., 2014). To examine the distribution of PGL condensates relative to MEG-3 condensates, we utilized the KT3 and OLLAS antibodies for immunostaining of untagged endogenous PGL-3 and OLLAS-tagged MEG-3. In embryos expressing wild-type MEG-3, MEG-3 and PGL-3 co-localize in posterior condensates that are segregated to the P₁ blastomere (Figure 3A). In embryos lacking meg-3 and meg-4, PGL-3 condensates distributed throughout the cytoplasm of the zygote and segregated equally to AB and P₁ (Figure 3A). We observed a similar pattern in embryos expressing MEG-3_IDR, MEG-3_698, and MEG-3_HMGL- indicating that none of these MEG-3 derivatives are sufficient to localize PGL condensates (Figure 3A, Figure 3—figure supplement 1A). In contrast, in embryos expressing MEG-3_Cterm, PGL-3 condensates preferentially assembled in P₁, although they were smaller and fewer than in wild-type (Figure 3A, B). Embryos expressing MEG-3_Cterm enriches PGL-3 in P₁, though not as efficiently as wild-type, while PGL-3 is not enriched in meg-3 meg-4, or embryos expressing MEG-3_IDR or MEG-3_HMGL- (Figure 3C). In wild-type 28-cell stage embryos, PGL-3 condensates are highly enriched in P₄. No such enrichment was observed in embryos expressing the MEG-3_Cterm or any other MEG-3 variant (Figure 3—figure supplement 1). We conclude that the MEG-3_Cterm is sufficient to enrich PGL-3 condensates in P blastomeres in early stages, but not sufficient to support robust PGL-3 localization through P₄.

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Enrichment of PGL-3 in P₁ in embryos expressing wild-type MEG-3 and variants.

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Wild-type MEG-3 condensates associate closely with the surface of PGL condensates (Putnam et al., 2019; Wang et al., 2014). With the resolution afforded by immunostaining, this configuration appears as co-localized MEG and PGL puncta in fixed embryos (Wang et al., 2014, Figure 3B). We found that PGL-3 condensates co-localized with MEG-3_Cterm condensates (37/37 PGL-3 condensates scored in P₁; Figure 3B) as in wild-type. In contrast, we observed no such co-localization with MEG-3_IDR or MEG-3_HMGL-. The MEG-3_IDR is mostly cytoplasmic and forms only rare condensates in P₁. We occasionally observed PGL condensates with an adjacent MEG-3_IDR condensate (5/19 PGL-3 condensates scored in P₁, Figure 3B), but these were not co-localized. Unlike the MEG-3_IDR, MEG-3_HMGL- forms many condensates in P₁, although these tended to be smaller than wild-type (Figure 3A, B). Still, although we occasionally observed PGL condensates with an adjacent MEG-3_HMGL- condensate (12/30 PGL-3 condensates scored in P₁; Figure 3B), we never observed fully overlapping PGL/MEG-3_HMGL- co-condensates. We conclude that, despite forming many condensates in P blastomeres, MEG-3_HMGL- condensates do not associate efficiently with, and do not support the localization of, PGL-3 condensates.

MEG-3 recruits mRNAs to P granules by direct binding that traps mRNA into the non-dynamic MEG-3 condensates (Lee et al., 2020). To determine which MEG-3 domain is required for mRNA recruitment to MEG-3 condensates in vivo, we performed in situ hybridization against the MEG-3-bound mRNA Y51F10.2. Prior to polarization, Y51F10.2 is uniformly distributed throughout the zygote cytoplasm (Figure 4A). Y51F10.2 becomes progressively enriched in P granules starting in the late one-cell stage and forms easily detectable micron-sized foci by the four-cell stage (Lee et al., 2020; Figure 4A). In contrast, in meg-3 meg-4 embryos, Y51F10.2 remains uniformly distributed in the cytoplasm at all stages. Strikingly, we observed the same failure to assemble Y51F10.2 foci in embryos expressing MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL- (Figure 4A). This was surprising since all three MEG-3 variants form visible condensates by the four-cell stage (Figure 3A).

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Y51F10.2 mRNA levels in embryos expressing wild-type MEG-3 and variants.

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To characterize the fate of Y51F10.2 transcripts in meg-3 meg-4 mutants, we compared the intensity of the Y51F10.2 in situ hybridization signal relative to a control RNA (T26A5.2) in one-cell and four-cell stage embryos (Figure 4B, Materials and methods, Figure 4—figure supplement 1). In wild-type, Y51F10.2 RNA levels do not change significantly from the one-cell to the four-cell stage. In contrast, in meg-3 meg-4 embryos, Y51F10.2 levels decreased by ~50% by the four-cell stage, despite starting at levels similar to wild-type in the one-cell stage. This finding is consistent with RNAseq results, which indicated lower levels of P granule mRNAs in meg-3 meg-4 embryos (Lee et al., 2020). We observed a similar loss of Y51F10.2 RNA in embryos expressing MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL- (Figure 4B). We repeated this analysis with a second MEG-3-bound mRNA, nos-2. Similar to Y51F10.2, nos-2 levels remained constant from the one-cell to four-cell stage in embryos expressing wild-type MEG-3, and decreased in meg-3 meg-4 embryos and embryos expressing MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL- (Figure 4—figure supplement 2). These results suggest that failure to recruit Y51F10.2 and nos-2 mRNAs into granules leads to their premature degradation.

After the four-cell stage, as has been reported for other maternal RNAs (Baugh et al., 2003; Seydoux and Fire, 1994), Y51F10.2 is rapidly turned over in somatic blastomeres. At the four-cell stage, Y51F10.2 mRNA levels were approximately twofold higher P₂ than in somatic blastomeres in wild-type embryos, and ~1.2-fold higher in meg-3 meg-4 embryos and in embryos expressing MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL- (Figure 4C). By the 28-cell stage, in wild-type embryos, Y51F10.2 levels were ~10-fold higher in the germline founder cell P₄ compared to somatic blastomeres. In contrast, in meg-3 meg-4 embryos, Y51F10.2 mRNA levels were only approximately twofold enriched over somatic levels. Similarly, in embryos expressing the MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL-, Y51F10.2 enrichment in P₄ averaged around approximately twofold (Figure 4A, D).

Enrichment of mRNAs in P granules can also be detected using an oligo-dT probe to detect polyadenylated mRNAs (Seydoux and Fire, 1994). In wild-type 28-cell stage embryos, a concentrated poly-A signal is detected around the nucleus of the P₄ blastomere. Poly-A signal intensity in P₄ is ~1.8-fold higher than that observed in somatic blastomeres (Figure 4—figure supplement 3A, B). This enrichment was not detected in meg-3 meg-4 mutants. In meg-3 meg-4 mutants, and in embryos expressing the three MEG-3 derivatives, poly-A signal intensity was similar between the somatic and P₄ blastomeres. The lack of poly-A enrichment in P₄ was particularly striking in the case of MEG-3_IDR and MEG-3_HMGL- since those variants assemble robust perinuclear condensates in P₄ (Figure 2A).

Failure to efficiently segregate and stabilize maternal mRNAs in P blastomeres has been linked to the partial penetrance maternal-effect sterility (~30%) of meg-3 meg-4 mutants (Lee et al., 2020). We observed similar levels of sterility in hermaphrodites derived from mothers expressing the MEG-3_IDR, MEG-3_Cterm, and MEG-3_HMGL- (Figure 4—figure supplement 3C). We conclude that the MEG-3 C-terminus, IDR, and HMG-like motif are all required for efficient mRNA recruitment to P granules, which in turn is required for enrichment and stabilization in the P lineage and robust germ cell fate specification.

We showed previously that MEG-3 readily condenses with RNA in vitro (Lee et al., 2020). To analyze the properties of MEG-3 variants in vitro, we expressed and purified His-tagged full-length MEG-3, MEG-3_Cterm, MEG-3_IDR, MEG-3_698, MEG-3_HMGL-, and MEG-3_{Cterm HMGL-}, a MEG-3_Cterm variant with a mutated HMGL motif (same alanine substitutions as in MEG-3_HMGL-). We first tested each variant for its ability to bind RNA. Using fluorescence polarization and gel shift assays, we previously showed that the MEG-3 IDR binds an RNA oligo (poly-U(30)) with near nanomolar affinity in vitro (Smith et al., 2016). We repeated these observations using a filter binding assay where proteins are immobilized on a filter to minimize possible interference due to condensation of MEG-3 in solution (Materials and methods, Figure 5A). Consistent with previous observations (Smith et al., 2016), we found that MEG-3_IDR and MEG-3₆₉₈ exhibit high affinity for poly-U(30) (Figure 5A, Figure 5—figure supplement 1B). Wild-type MEG-3 also bound poly-U(30) efficiently albeit at a lower affinity compared to MEG-3_IDR and MEG-3₆₉₈. In contrast, MEG-3_Cterm exhibited negligible RNA binding (Figure 5A). HMG domains are common in DNA-binding proteins and have been shown to mediate protein:nucleic acid interactions in vivo (Genzor and Bortvin, 2015; Reeves, 2001; Thapar, 2015). MEG-3_HMGL- bound to RNA as efficiently as full-length MEG-3, indicating that the HMGL motif does not contribute to RNA binding in MEG-3 (Figure 5A).

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RNA binding of MEG-3 and variants in vitro.

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In vivo, MEG-3 binds to maternal mRNAs including nos-2 (Lee et al., 2020). To examine the affinity of MEG-3 for nos-2 RNA, we repeated the filter binding assay with labeled poly-U(30) using unlabeled nos-2 RNA as a competitor (Figure 5B). The competition assay revealed that full-length MEG-3 and MEG-3_IDR bind to nos-2 RNA with the same high affinity. We conclude that the MEG-3 IDR is necessary and sufficient for RNA binding.

To determine which regions of MEG-3 are required for condensation in vitro, MEG-3 and variants were trace-labeled with covalently attached fluorophores and examined for condensate formation by microscopy. MEG-3 condensation is sensitive to salt and RNA concentration with RNA having a strong solubilizing influence especially in low salt (Lee et al., 2020). We tested four conditions varying RNA and salt and keeping MEG-3 concentration constant at 150 nM near the physiological range (Figure 6A, Figure 6—figure supplement 1). At that concentration, MEG-3 and variants were all soluble in low salt/high RNA condensation buffer (50 mM NaCl to 80 ng/μL nos-2 RNA) (Figure 6—figure supplement 1). Under higher salt conditions (150 mM NaCl salt and 20 ng/μL or 80 ng/μL nos-2 RNA), MEG-3_IDR and MEG-3₆₉₈ remained mostly soluble forming only rare condensates (Figure 6A, Figure 6—figure supplements 1, 2). In contrast, full-length MEG-3 formed robust condensates under those conditions. MEG-3_C-term also formed condensates, albeit with lower efficiency compared to MEG-3 (Figure 6A, Figure 6—figure supplement 1). The MEG-3_Cterm condensates were also approximately twofold less efficient at recruiting RNA compared to full-length MEG-3 and MEG-3_IDR (Figure 6A, Figure 6—figure supplement 1B). We conclude that, as observed in vivo, the C-terminus is the primary driver of MEG-3 condensation. The MEG-3 C-terminus is not sufficient, however, for maximum condensation or RNA recruitment, which additionally require the IDR.

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MEG-3 and variants condensation with and without PGL-3 in vitro.

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Surprisingly, under all conditions, MEG-3_HMGL- and MEG-3_CtermHMGL- behaved identically to MEG-3 and MEG-3_Cterm, respectively (Figure 6, Figure 6—figure supplement 1), indicating that the HMGL motif is dispensable for RNA recruitment and condensation in vitro. HMGL was required for RNA recruitment and efficient condensation in vivo (Figure 4), suggesting that our in vitro conditions do not fully reproduce in vivo conditions (see below).

When combined in condensation buffer, MEG-3 and PGL-3 form co-condensates that resemble the architecture of P granules in vivo, with the smaller MEG-3 condensates (~100 nm) forming a dense layer on the surface of the larger PGL-3 condensates (Putnam et al., 2019; Figure 6B, Figure 6—figure supplement 3A). We found that the MEG-3_Cterm formed co-condensates with PGL-3 that were indistinguishable from those formed by wild-type MEG-3 (Figure 6—figure supplement 3A, B). The MEG-3_IDR, in contrast, mixed homogenously with the PGL-3 phase as previously reported (Putnam et al., 2019, Figure 6—figure supplement 3A). MEG-3_HMGL- and MEG-3_CtermHMGL- behaved as MEG-3 and MEG-3_Cterm, respectively. In vivo, MEG-3_HMGL- does not associate efficiently with PGL-3 condensates (Figure 3B), again suggesting that in vitro conditions do not reproduce the more stringent in vivo environment.

We repeated the co-condensation assays in the absence of RNA using a higher concentration of PGL to force PGL condensation in the absence of RNA. Under these conditions, MEG-3 and PGL-3 formed co-condensates that were indistinguishable from co-condensates assembled in the presence of RNA (compare right and left panels, Figure 6—figure supplement 3A and B). Again, the C-terminus was necessary and sufficient for co-assembly. MEG-3_IDR homogenously mixed with the PGL-3 phase and did not form independent condensates. We conclude that condensation of MEG-3 on the surface of PGL condensates requires the MEG-3 C-terminus and does not require RNA in vitro.

To examine the ability of MEG-3 to recruit nos-2 RNAs to MEG-3/PGL-3 co-condensates, we assembled the co-condensates in the presence of labeled 40 ng/μL labeled nos-2 RNA and unlabeled 150 μM poly-U(30). Unlabeled poly-U(30) was necessary to prevent nos-2 RNA from accumulating non-specifically in the PGL-3 phase. Under these high RNA conditions, all variants were recruited to PGL condensates, but only full-length MEG-3 (and MEG-3_HMGL-) formed a distinctive layer around PGL condensates that recruited RNA above the background level recruited to the PGL-3 phase (Figure 6B–D). MEG-3_IDR and MEG-3_Cterm, in contrast, mixed with the PGL phase and did not enrich nos-2 RNA above background levels (Figure 6B, C). These observations suggest that enrichment of nos-2 RNA in MEG-3/PGL-3 co-condensates requires robust MEG-3 condensation driven by the combined action of the MEG-3 IDR and C-terminus.

We reported previously that MEG-3 binds directly to PGL-1, as determined in a GST-pull-down assay using partially purified recombinant proteins (Wang et al., 2014). HMG domains have been implicated in protein-protein interactions (Reeves, 2001; Stros et al., 2007; Wilson and Koopman, 2002). To examine whether the HMG domain is required to mediate MEG-3/PGL binding, we repeated the GST-pull-down assay using fusion proteins of GST::MEG-3_Cterm and MBP::PGL-1 and PGL-3 (Figure 7A). GST::MEG-3_IDR fusions were not expressed and thus could not be tested in this assay. We found that the GST::MEG-3_Cterm binds efficiently to PGL-1 and PGL-3, but not to MBP or to an unrelated control protein PAA-1 (Figure 7A, B). A GST::MEG-3_Cterm fusion with mutations in the HMG-like domain bound less efficiently to both PGL-1 and PGL-3 (Figure 7A, B). To complement the GST assay, we examined the ability of purified labeled PGL-3 to bind to beads coated with purified labeled MEG-3 derivatives (bead halo assay) (Figure 7C, Figure 7—figure supplement 1A-C). We found that PGL-3 is recruited efficiently to beads coated with MEG-3_C-term, but not to beads coated with MEG-3_IDR or MEG-3_{Cterm HMGL-} (Figure 7C, Figure 7—figure supplement 1A). We conclude that the MEG-3_C-term binds directly to PGL-3 and that this interaction requires the HMGL motif. This finding provides a potential explanation for why the HMGL is required for co-assembly of MEG-3 and PGL-3 condensates in vivo but not in vitro. Direct binding between MEG-3 and PGL-3 molecules may be necessary to assemble co-condensates in the crowded in vivo environment and may not be required in vitro where no other proteins or condensates compete for binding to MEG-3 or PGL-3.

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MEG-3 and variants binding to PGL-3.

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The MEG-3 C-terminus contains an HMG-like motif required to bind to PGL-3 and additional predicted low-disorder sequences that drive condensation by an unknown mechanism. The HMGL domain is not required for condensation in vitro but is required for maximal condensation efficiency in vivo. We suggest the HMGL domain enhances condensation indirectly in vivo by concentrating MEG-3 on the surface of PGL droplets, which stimulates condensation (Putnam et al., 2019). Docking of P bodies on stress granules has been proposed to involve RNA:RNA duplexes (Tauber et al., 2020). In contrast, we find that docking of MEG-3 condensates on PGL condensates does not require RNA in vitro and can occur in the absence of any visible RNA enrichment in vivo. Most strikingly, mutations in the HMGL domain that prevent binding between MEG-3 and PGL-3 molecules in solution prevent docking of MEG-3 and PGL condensates in vivo. Together, these observations suggest that MEG-3 condensation and co-assembly with PGL-3 condensates is driven primarily by protein-protein interactions and does not require RNA. We note that the HMGL domain is dispensable for co-assembly of MEG-3 and PGL-3 condensates in vitro, indicating that our condensation assay conditions do not fully reproduce the stringent environment of the cytoplasm.

We previously reported that enrichment of PGL droplets to the posterior of the zygote requires meg-3 (Smith et al., 2016; Wang et al., 2014). Our new findings suggest that this activity is linked to MEG-3’s ability to associate stably with the PGL interface. MEG-3_Cterm, which is sufficient for MEG-3/PGL co-assembly, is sufficient to localize PGL in zygotes. Conversely, MEG-3_HMGL- condensates, which do not interact stably with PGL condensates, fail to enrich PGL condensates in the posterior of zygotes. PGL localization involves preferential growth and dissolution of PGL droplets in the anterior and posterior, respectively. One possibility is that tight binding of MEG condensates lowers the surface tension of PGL droplets allowing MEG/PGL co-assemblies in the posterior to grow at the expense of the less stable, ‘naked’ PGL droplets in the anterior.

What enriches MEG-3 condensates in the posterior? We previously hypothesized that MEG-3 asymmetry is driven by a competition for RNA between the MEG-3 IDR and MEX-5, an RNA-binding protein that acts as an RNA sink in the anterior (Smith et al., 2016). Consistent with this hypothesis, the MEG-3 IDR is sufficient to enrich MEG-3 in posterior cytoplasm. Unexpectedly, however, we found that the MEG-3_Cterm condenses preferentially in the zygote posterior despite uniform distribution in the cytoplasm, suggesting that additional mechanisms acting on the MEG-3 C-terminus contribute to MEG-3 regulation in space. Consistent with this view, a recent study examining MEG-3 dynamics by single-molecule imaging (Wu et al., 2019) found that the slowly diffusing MEG-3 molecules that populate the MEG-3 gradient in the cytoplasm represent a distinct population of MEG-3 molecules from those that associate with PGL droplets. We propose that MEG-3 asymmetry is sustained by two independent mechanisms: one acting on the MEG-3 C-terminus that biases condensation of MEG-3 in posterior cytoplasm and one acting on the MEG-3 IDR that enriches MEG-3 molecules in posterior cytoplasm. We speculate that the latter may serve to segregate high levels of MEG-3 to P blastomeres needed to support PGL asymmetry through the P₄ stage. Consistent with this view, the MEG-3_Cterm, which does not enrich in a gradient, is not sufficient to localize PGL condensates in P blastomeres past the four-cell stage.

The MEG-3 IDR binds RNA with high-affinity in vitro but is not sufficient to enrich RNA in vivo despite forming some condensates. RNA recruitment in vivo additionally requires the MEG-3 C-terminus including the HMG-like motif. These observations suggest that maximal MEG-3 condensation is essential to build a protein scaffold that can support stable RNA recruitment in vivo. Separate domains for RNA-binding and protein condensation have also been observed for other germ granule scaffolds. For example, the Balbiani body protein Xvelo uses a prion-like domain to aggregate and a separate RNA-binding domain to recruit RNA (Boke et al., 2016). Similarly, condensation of Drosophila Oskar does not require the predicted Oskar RNA-binding domain, although this domain augments condensation (Kistler et al., 2018). These observations parallel our findings with MEG-3 and contrast with recent findings reported for the stress granule scaffold G3BP. Condensation of G3BP in vitro requires RNA and two C-terminal RNA-binding domains. A N-terminal dimerization domain is also required but, unlike the prion-like domain of Xvelo or the C-terminus of MEG-3, is not sufficient to drive condensation on its own. Dimerization of G3BP is thought to enhance LLPS indirectly by augmenting the RNA-binding valency of G3BP complexes. G3BP also contains an inhibitory domain that gates its RNA-binding activity and condensation at low RNA concentrations. This modular organization ensures that G3BP functions as a sensitive switch that initiates LLPS when sufficient RNA molecules are available to cross-link G3BP dimers into a large network (Guillén-Boixet et al., 2020; Yang et al., 2020). Stress granules are transient structures that form under conditions of general translational arrest where thousands of transcripts are released from ribosomes. In contrast, germ granules are long-lived structures that assemble in translationally active cytoplasm and recruit only a few hundred specific transcripts (~500 in C. elegans embryos) (Jamieson-Lucy and Mullins, 2019; Lee et al., 2020; Trcek and Lehmann, 2019; Updike and Strome, 2010). One possibility is that protein-based condensation mechanisms may be better suited to assemble long-lived granules able to capture and retain rare transcripts. By concentrating IDRs with affinity for RNA, protein scaffolds could act as seeds for localized LLPS to amplify protein and RNA condensation. Consistent with this view, IDRs have been observed to undergo spontaneous LLPS in cells when artificially tethered to protein modules that self-assemble into large multimeric structures (Nakamura et al., 2019). A challenge for the future will be to understand the mechanisms that regulate the assembly and disassembly of protein scaffolds at the core of germ granules.

C. elegans was cultured at 20° C according to standard methods (Brenner, 1974). To measure maternal-effect sterility, 10 gravid adults were picked to an OP50 plate and allowed to lay eggs for ~2 hr, then removed. Adult progeny were scored for empty uteri (white sterile phenotype) under a dissecting microscope.

MEG-3 and MEG-4 protein sequences were aligned with HMG boxes from GCNA proteins of Caenorhabditis and example vertebrates along with the canonical HMG box of mouse SOX3 using MUSCLE (Edgar, 2004). Alignment was manually adjusted according to the published CGNA HMG Hidden Markov Model (Carmell et al., 2016). Amino acids were chosen for mutation based on conservation in nematodes.

Genome editing was performed in C. elegans using CRISPR/Cas9 as described in Paix et al., 2017. Strains used in this study, along with guides and repair templates, are listed in Supplementary file 1. Some strains were generated in two steps. For example, MEG-3_HMGL- was generated by deleting the entire HMGL-like motif in a first step (JH3632) and inserting a modified HMG-like motif with the desired mutations in a second step (JH3861). Genome alterations were confirmed by Sanger sequencing, and expression of tagged strains was verified by immunostaining and western blotting (Figure 2—figure supplement 1C).

On all scatterplots, central bars indicate the mean and error bars indicate one standard deviation. Unless otherwise indicated, differences within three or more groups were evaluated using a one-factor ANOVA and differences between two groups using an unpaired Student’s t-test.

Fluorescence confocal microscopy for Figure 2—figure supplement 1A and Figure 6—figure supplements 1, 2 was performed using a Zeiss Axio Imager with a Yokogawa spinning-disc confocal scanner. Fluorescence confocal microscopy for all other figures was performed using a custom-built inverted Zeiss Axio Observer with CSU-W1 Sora spinning disk scan head (Yokogawa), 1×/2.8× relay lens (Yokogawa), fast piezo z-drive (Applied Scientific Instrumentation), and a iXon Life 888 EMCCD camera (Andor). Samples were illuminated with 405/488/561/637 nm solid-state laser (Coherent), using a 405/488/561/640 transmitting dichroic (Semrock) and 624-40/692-40/525-30/445-45 nm bandpass filter (Semrock), respectively. Images from either microscope were taken with using Slidebook v6.0 software (Intelligent Imaging Innovations) using a 40×–1.3 NA/63×–1.4 NA objective (Zeiss) depending on sample.

Adult worms were placed into M9 on poly-l-lysine (0.01%)-coated slides and squashed with a coverslip to extrude embryos. Slides were frozen by laying on aluminum blocks pre-chilled with dry ice for >5 min. Embryos were permeabilized by freeze-cracking (removal of coverslips from slides) followed by incubation in methanol at −20°C for >15 min and in acetone −20°C for 10 min. Slides were blocked in PBS-Tween (0.1%) BSA (0.5%) for 30 min at room temperature and incubated with 50 μL primary antibody overnight at 4°C in a humid chamber. For co-staining experiments, antibodies were applied sequentially (OLLAS before KT3, K76) to avoid cross-reaction. Antibody dilutions (in PBST/BSA): KT3 (1:10, DSHB), K76 (1:10 DSHB), and Rat αOLLAS-L2 (1:200, Novus Biological Littleton, CO), Secondary antibodies were applied for 2 hr at room temperature. Samples were mounted Prolong Diamond Antifade Mountant or VECTASHIELD Antifade Mounting Media with DAPI. Embryos were staged using DAPI stained nuclei and 25 confocal slices spaced 0.18 µm apart and centered on the P cell nucleus were taken using a 63× objective. Unless otherwise indicated, images presented in figures are maximum projections.

All analyses were performed in ImageJ. For measurements of embryos/cells (Figure 2B-D, 3C), confocal stacks were sum projected and the integrated density was measured within a region of interest. For measurements of condensate intensity and number (Figure 2B-D), the 3D objects' counter function was used with a minimum size of 10 pixels on the full confocal stack confined to a region of interest drawn around the P cell and including objects on edges. The integrated density for all identified particles was summed to give the total intensity in condensates.

smFISH probes were designed using Biosearch Technologies’s Stellaris Probe Designer, with the fluorophor Quasar670. For sample preparation, embryos were extruded from adults on poly-l-lysine (0.01%) slides and subjected to freeze-crack followed by methanol fixation at >20°C for >15 min. Samples were washed five times in PBS-Tween (0.1%) and fixed in 4% PFA (Electron Microscopy Science, No. 15714) in PBS for 1 hr at room temperature. Samples were again washed four times in PBS-Tween (0.1%), twice in 2× SCC, and once in wash buffer (10% formamide, 2× SCC) before blocking in hybridization buffer (10% formamide, 2× SCC, 200 µg/mL BSA, 2 mM Ribonucleoside Vanadyl Complex, 0.2 mg/mL yeast total RNA, 10% dextran sulfate) for >30 min at 37°C. Hybridization was then conducted by incubating samples with 50 nM probe solutions in hybridization buffer overnight at 37°C in a humid chamber. Following hybridization, samples were washed twice in wash buffer at 37°C, twice in 2× SCC, once in PBS-Tween (0.1%) and twice in PBS. Samples were mounted Prolong Diamond Antifade Mountant.

All measurements were performed on a single confocal slice centered on the P cell nucleus in ImageJ. For early embryos where there is distinct punctate signal (one- and four-cell stage; Figure 4B, C, Figure 4—figure supplements 1,2), a region of interest was drawn, the Analyze Particles feature was used with a manual threshold to identify and measure the integrated density of the puncta. The raw integrated density for all particles in the region of interest was summed to give the total intensity of the mRNA in that region. For 28-cell embryos (Figure 4D, Figure 4—figure supplement 3), a region of interest was drawn around the P₄ blastomere and the intensity of that region was divided by the intensity of a region of the same size in the anterior soma.

Worms were synchronized by bleaching to collect embryos, shaken approximately 20 hr in M9, then plating on large enriched peptone plates with a lawn of Escherichia coli NA22 bacteria. Embryos were harvested from young adults (66 hr after starved L1 plating) and sonicated in 2% SDS, 65 mM Tris pH 7, 10% glycerol with protease and phosphatase inhibitors. Lysates were spun at 14,000 rpm for 30 min at 4°C and cleared supernatants were transferred to fresh tubes. Lysates were run on 4–12% Bis-Tris pre-cast gels (Bio-Rad Hercules, CA). Western blot transfer was performed for 1 hr at 4°C onto PVDF membranes. Membranes were blocked overnight and washed in 5% milk, 0.1% Tween-20 in PBS; primary antibodies were incubated overnight at 4°C; secondary antibodies were incubated for 2 hr at room temperature. Membranes were first probed for OLLAS then stripped by incubating in 62.5 mM Tris HCl pH 6.8, 2% SDS, 100 mM ß-mercaptoethanol at 42°C. Membranes were then washed, blocked, and probed for α-tubulin. Antibody dilutions in 5% milk/PBST: Rat α OLLAS-L2 (1:1000, Novus Biological Littleton, CO), Mouse α-tubulin (1:1000, Sigma, St. Louis, MO).

mRNAs were transcribed using T7 mMessageMachine (Thermo Fisher) using the manufacturer’s recommendation as described (Lee et al., 2020). Fluorescently labeled mRNAs were generated by including a trace amount of ChromaTide Alexa Fluor 488–5-UTP or 546–14-UTP in the transcription reaction. Template DNA for transcription reactions was obtained by PCR amplification from plasmids. Free NTPs and protein were removed by lithium chloride precipitation. RNAs were resuspended in water and stored at −20°C. The integrity of RNA products was verified by agarose gel electrophoresis.

Protein condensation was induced by diluting proteins out of storage buffer into condensation buffer containing 25 mM HEPES (pH 7.4), salt adjusted to a final concentration of 50 or 150 mM (37.5 mM KCl, 12.5 or 112.5 mM NaCl), and RNA. For MEG-3 and PGL-3 co-condensate experiments with RNA, we used 150 nM MEG-3, 1.8 μM PGL-3. For MEG-3 and PGL-3 co-condensate poly-U(30) competition experiments with nos-2, we used 150 nM MEG-3, 2.5 μM PGL-3. For co-assembly experiments in the absence of RNA, we used 150 nM MEG-3, 5 μM PGL-3. MEG-3 and PGL-3 solutions contained 10–50 nM fluorescent trace labels with either 488 or 647 (indicated in the figure legends). MEG-3 and PGL-3 condensation reactions with nos-2 and poly-U(30) RNA were incubated at room temperature for 30 min before spotting onto a No. 1.5 glass bottom dish (Mattek) and imaged using a 40× with a 1× (used for quantification) and 2.8× (used for display) relay lens oil objective (Figure 6B). All other condensate reactions were imaged using thin-chambered glass slides (Erie Scientific Company 30-2066A) with a coverslip (Figure 6—figure supplements 1, 2). Images are single planes acquired using a 40× oil objective over an area spanning 171 × 171 μm.

To quantify the relative intensity of MEG-3 in condensates, a mask was created by thresholding images, filtering out objects of less than four pixels to minimize noise, applying a watershed filter to improve separation of objects close in proximity, and converting to a binary image by the Otsu method using the nucleus counter cookbook plugin. Minimum thresholds were set to the mean intensity of the background signal of the image plus 1–2 standard deviations. The maximum threshold was calculated by adding 3–4 times the standard deviation of the background. Using generated masks, the integrated intensity within each object was calculated. Any normalization is indicated in the corresponding figure legend. Each replicate contained three images each spanning an area of 171 × 171 μm (Figure 6A, Figure 6—figure supplement 1B and C) or 316.95 × 316.95 μm (Figure 6C, D).

Proteins were step-dialyzed from 6 M urea into 4.5 M urea, 3 M urea, 1.5 M urea, and 0 M urea in MEG-3 storage buffer (25 mM HEPES, pH 7.4, 1 M NaCl, 6 mM ß-mercaptoethanol, 10% glycerol). RNA-binding reactions consisted of 50 nM 3′ fluorescein-labeled 30U RNA oligonucleotides (poly-U(30)) incubated with either varying protein concentrations (direct binding of polyU(30)) or constant concentrations of protein and varying concentrations of nos-2 mRNA (long RNA binding by competition) for 30 min at room temperature (final reaction conditions 3.75 mM HEPES, 150 mM NaCl, 0.9 mM, 0.9 mM ß-mercaptoethanol, 1.5% glycerol, 10 mM Tris HCl). Fluorescence filter binding protocol was adapted from a similar protocol using radiolabeled RNA (Rio, 2012). Briefly, a pre-wet nitrocellulose was placed on top of Hybond-N+ membrane in a dot-blot apparatus, reactions were applied to the membranes, then washed 2× with 10 mM Tris HCl. Membranes were briefly dried in air, then imaged using a typhoon FLA-9500 with blue laser at 473 nm. Fraction of RNA bound for each reaction was calculated by dividing the fluorescence signal on the nitrocellulose membrane by the total signal from both membranes.

For direct binding (Figure 5A), K_d was calculated by plotting the bound fraction of poly-U(30) RNA as a function of protein concentration and fitting to the following equation in Prism8, where P is the protein concentration in nM, B is the bound fraction of poly-U(30) with non-specific binding subtracted, B_max is the maximum specific binding, K_d is the concentration needed to achieve a half-maximum binding at equilibrium, and h is the Hill slope:

For competition binding of nos-2 RNA (Figure 5B), IC50 was calculated by plotting the bound fraction of poly-U(30)as a function of the log of the concentration of nos-2 and fitting to the following equation, where B is the bound fraction of poly-U(30) in nM, B_max is the maximum fraction bound, B_min is the minimum fraction bound, X is the concentration of nos-2 RNA (in nM 30mers), and IC50 is the concentration of nos-2 RNA (in nM 30mers) needed to achieve half-maximum inhibition of poly-U(30) binding:

For a more direct comparison between MEG-3 variants that have a different K_d, the EC50 was converted to K_i using the following equation, where [30U] is the concentration of fluorescent labeled poly-U(30) in the reaction in nM and K_d is the experimentally determined K_d for that MEG-3 variant and poly-U(30):

All binding constants are the average value from fitting each replicate separately.

GST fusion proteins were cloned into pGEX6p1 (GE Healthcare, Pittsburgh, PA). MBP fusion proteins were cloned into pJP1.09, a Gateway-compatible pMAL-c2x (Pellettieri et al., 2003). Proteins were expressed in Rosetta E. coli BL21 cells grown for approximately 4 hr at 37°C, then induced with 1 mM IPTG and grown overnight at 16°C. 200 mg of bacterial pellet of GST fusion proteins was resuspended in 50 mM HEPES, 1 mM EGTA, 1 mM MgCl₂, 500 mM KCl, 0.05% NP40, 10% glycerol, pH 7.4 (IP Buffer) with protease and phosphatase inhibitors, lysed by sonication, and bound to magnetic GST beads. Beads were washed and incubated with MBP fusion proteins at 4°C for 1 hr in the same buffer as for lysis. After washing, beads were eluted by boiling and eluates were loaded on SDS-PAGE. Western blot transfer was performed for 1 hr at 4°C onto PVDF membranes. Membranes were blocked and washed in 5% milk, 0.1% Tween-20 in PBS, and incubated with HRP conjugate antibodies. Antibody dilutions in 5% milk/PBST:anti-MBP HRP conjugated, 1:50,000 (NEB, and anti-GST HRP conjugates, 1:2000) (GE Healthcare). Scanned western blot films were quantified using the gel analysis tool in ImageJ.

Fluorescent protein bead halo assay was adapted from Patel and Rexach, 2008. 50 μL of Nickel-NTA agarose beads (Qiagen) were incubated with 50 μL of 10 μM MEG-3 derivatives trace-labeled with Alexa647 or no protein in MEG-3 storage buffer for 1 hr. Beads were washed five times with IP Buffer then blocked for 1 hr in blocking buffer (4 mg/mL BSA, 50 mM HEPES, 1 mM EGTA, 1 mM MgCl₂, 500 mM KCl, 0.05% NP40, 10% glycerol, pH 7.4). MBP and PGL-3 trace-labeled with Alexa555 were prepared in PGL-3 storage buffer and diluted to 3 μM in blocking buffer. Additional concentrations of PGL-3 were diluted from this solution to maintain the ratio of label to total protein. 5 μL of blocked MEG-3 or empty beads was added to 50 μL of PGL-3 or MBP solution and incubated for 1 hr. Beads were washed five times in blocking buffer and resuspended in PBS. Beads were spotted onto a No. 1.5 glass-bottom dish (Mattek) and imaged using a 10× air objective. Images are single planes through the center of the beads.

To quantify the relative intensity of PGL-3 to MEG-3 derivatives on the beads, a mask was created using the Analyze Particles feature in ImageJ on the MEG-3 channel, using a minimum size of 10⁻⁵ cm². Using the generated mask, the integrated intensity within each bead was calculated for both the MEG-3 and MBP/PGL-3 channels. To remove non-specific binding signal, the mean intensity empty beads incubated with MBP or PGL-3 were subtracted from each pixel yielding the total intensity of each bead. To calculate the intensity ratio for each image, the total intensity on beads of MBP or PGL-3 was divided by the total intensity of MEG-3 on beads. This ratio was normalized to the mean MBP ratio.

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

We thank the Johns Hopkins Neuroscience Research Multiphoton Imaging Core (NS050274) and the Johns Hopkins Integrated Imaging Center (S10OD023548) for excellent microscopy support. We thank the Page lab for assistance identifying the HMG-like motif, the Nathans lab for the OLLAS antibody, and Addgene for TEV protease from the Waugh lab. We thank Deepika Calidas for collaboration on strains JH3479, JH3517, and JH3420, Mario Martinez for assistance in protein purification, and Baltimore Worm Club and the Seydoux lab for many helpful discussions. This work was supported by the National Institutes of Health (grant number GS: 5R37HD037047, AP: F32GM134630). GS is an investigator of the Howard Hughes Medical Institute.

© 2021, Schmidt et al.

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