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HDAC1 SUMOylation promotes Argonaute-directed transcriptional silencing in C. elegans

  1. Heesun Kim
  2. Yue-He Ding
  3. Gangming Zhang
  4. Yong-Hong Yan
  5. Darryl Conte Jr
  6. Meng-Qiu Dong
  7. Craig C Mello  Is a corresponding author
  1. RNA Therapeutics Institute, University of Massachusetts Medical School, United States
  2. National Institute of Biological Sciences, China
  3. Howard Hughes Medical Institute, United States
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Cite this article as: eLife 2021;10:e63299 doi: 10.7554/eLife.63299

Abstract

Eukaryotic cells use guided search to coordinately control dispersed genetic elements. Argonaute proteins and their small RNA cofactors engage nascent RNAs and chromatin-associated proteins to direct transcriptional silencing. The small ubiquitin-like modifier (SUMO) has been shown to promote the formation and maintenance of silent chromatin (called heterochromatin) in yeast, plants, and animals. Here, we show that Argonaute-directed transcriptional silencing in Caenorhabditis elegans requires SUMOylation of the type 1 histone deacetylase HDA-1. Our findings suggest how SUMOylation promotes the association of HDAC1 with chromatin remodeling factors and with a nuclear Argonaute to initiate de novo heterochromatin silencing.

Introduction

Argonautes are an ancient family of proteins that utilize short nucleic acid guides (usually composed of 20–30 nts of RNA) to find and regulate cognate RNAs (reviewed in Meister, 2013). Argonaute-dependent small RNA pathways are linked to chromatin-mediated gene regulation in diverse eukaryotes, including plants, protozoans, fungi, and animals (reviewed in Martienssen and Moazed, 2015). Connections between Argonautes and chromatin are best understood from studies in fission yeast Schizosaccharomyces pombe, where the Argonaute, Ago1, a novel protein, Tas3, and a heterochromatin protein 1 (HP1) homolog, Chp1, comprise an RNA-induced transcriptional silencing (RITS) complex that maintains and expands heterochromatin (Verdel et al., 2004). The Chp1 protein binds H3K9me3 through its conserved chromodomain (Partridge et al., 2000; Partridge et al., 2002) and is thought to anchor Ago1 within heterochromatin, where it is poised to engage nascent RNA transcripts (Holoch and Moazed, 2015). Low-level transcription of heterochromatin is thought to create a platform for propagating small-RNA amplification and heterochromatin maintenance (reviewed in Holoch and Moazed, 2015).

In yeast, a protein complex termed SHREC (Snf2/Hdac-containing Repressor complex) has been linked to both the establishment and maintenance of heterochromatin and transcriptional silencing (Job et al., 2016; Motamedi et al., 2008; Sugiyama et al., 2007). SHREC contains a homolog of type 1 histone deacetylase (HDAC), a homolog of Mi-2 and CHD3 ATP-dependent chromatin remodelers, and a Krüppel-type C2H2 zinc finger protein. SHREC therefore resembles the nucleosome remodeling and deacetylase (NuRD) complex in animals (Denslow and Wade, 2007; Torchy et al., 2015). NuRD complexes play a key role in converting chromatin from an active to a silent state, and can be recruited to targets through sequence-independent interactions (e.g., modified chromatin or methylated DNA) or through sequence-specific interactions via the Krüppel-type C2H2 zinc finger protein and other DNA-binding factors (Ecco et al., 2017; Lupo et al., 2013). In animals, NuRD complexes function broadly in developmental gene regulation and transposon silencing (Ecco et al., 2017; Feschotte and Gilbert, 2012; Ho and Crabtree, 2010).

The post-translational modification of heterochromatin factors by the small ubiquitin-like protein SUMO has been implicated at several steps in the establishment and maintenance of transcriptional gene silencing and has been linked to silencing mediated by the Piwi Argonaute (reviewed in Ninova et al., 2019). The addition of SUMO (i.e., SUMOylation) requires a highly conserved E2 SUMO-conjugating enzyme, UBC9, which interacts with substrate-specific co-factor (E3) enzymes to covalently attach SUMO to lysines in target proteins (Johnson, 2004). Whereas ubiquitylation is primarily associated with the protein turnover (reviewed in Glickman and Ciechanover, 2002), SUMOylation is primarily associated with changes in protein interactions, especially with proteins that contain SUMO-interacting motifs (SIMs) (reviewed in Kerscher, 2007). In mammals, for example, SUMOylation of the KAP1 corepressor is required to recruit the NuRD complex and the SETDB histone methyltransferase via SIM domains in CHD3 and SETDB and to silence KRAB targets (Ivanov et al., 2007).

Here, we identify a connection between the SUMO pathway and transcriptional silencing initiated by the Piwi Argonaute pathway in the Caenorhabditis elegans germline. We show that SUMOylation of C-terminal lysines on the type 1 HDAC, HDA-1, is required for Piwi-mediated transcriptional silencing. SUMOylation of HDA-1 promotes its association with conserved components of the C. elegans NuRD complex, the nuclear Argonaute HRDE-1/WAGO-9, the histone demethylase SPR-5, and the SetDB-related histone methyltransferase MET-2. Our findings suggest how SUMOylation of HDAC1 promotes the recruitment and assembly of an Argonaute-guided chromatin remodeling complex that orchestrates de novo transcriptional gene silencing in the C. elegans germline.

Results

The SUMO and HDAC pathways promote piRNA silencing

In C. elegans, silencing initiated by the Piwi Argonaute PRG-1 depends on chromatin modifications at the target locus and on a group of worm-specific Argonautes (WAGOs), including nuclear-localized family members WAGO-9/HRDE-1 and WAGO-10 (Ashe et al., 2012; Bagijn et al., 2012; Lee et al., 2012; Shirayama et al., 2012) and nuage-localized family members WAGO-1 and WAGO-4 (Gu et al., 2009; Shirayama et al., 2012; Xu et al., 2018). WAGOs engage antisense guides produced by cellular RNA-dependent RNA polymerases (RdRPs) (Gu et al., 2009). How the downstream machinery that amplifies and maintains silencing is recruited to targets remains unknown. To identify additional components of the transcriptional silencing arm of the piRNA pathway, we performed an RNAi-based genetic screen of chromatin factors and modifiers using a sensor transgene silenced by the piRNA pathway (Figure 1A; see Seth et al., 2018). The piRNA sensor is 100% silenced in wild-type germlines, but is desilenced in the germlines of prg-1(tm872), rde-3(ne3370), and hrde-1/wago-9(ne4769) mutants, resulting in expression of a bright, easily scored GFP::CSR-1 fusion protein (Figure 1BSeth et al., 2018). Even the partial inactivation of known piRNA silencing factors activated sensor expression in a percentage of exposed animals (Figure 1C and Supplementary file 1).

SUMOylation and chromatin remodeling factors promote piRNA-mediated silencing.

(A) Schematic of the piRNA sensor screen. The piRNA sensor strain contains a gfp::csr-1 transgene that is silenced by the piRNA pathway in the presence of an active oma-1::gfp transgene (Seth et al., 2018). OMA-1::GFP localizes to the cytoplasm of oocytes. Inactivation of the piRNA pathway (by RNAi, mutation, or auxin-inducible protein depletion) desilences the transgene, resulting in GFP::CSR-1 expression in perinuclear P-granules throughout the germline, as shown in (B). (B) Differential interference contrast and epifluorescence images of dissected gonads in wild-type (wt), prg-1(tm872), and wago-9/hrde-1(ne4769) worms. PRG-1 is required to initiate silencing, while WAGO-9 is required to maintain silencing. The percentage of desilenced worms and number of worms scored are shown. (C) Analysis of SUMO and chromatin remodeling factors required for piRNA-mediated silencing. Genes identified in the RNAi-based screen of chromatin factors are listed with their human homologs and with the percentage of worms that express GFP::CSR-1 among the total number of worms analyzed (n) when function is reduced by RNAi (blue column) or by either mutation or degron-dependent protein depletion (peach column).

Our RNAi screen identified many components of known HDAC complexes, as well as SUMO pathway factors (Figure 1C and Supplementary file 1). For example, depletion of mep-1 (Krüppel-type zinc finger protein) and other genes encoding NuRD-complex co-factors (let-418/Mi-2, hda-1/HDAC1, lin-40/MTA2/3, lin-53/RBBP4/7, and dcp-66/GATAD2B) desilenced the piRNA sensor (Figure 1C and Supplementary file 1). Depletion of SIN3-HDAC complex genes, sin-3 (SIN3) and mrg-1 (MORF4L1), also desilenced the reporter (Figure 1C and Supplementary file 1). RNAi of two SUMO pathway genes, smo-1 (SUMO) and ubc-9 (SUMO-conjugating enzyme), desilenced the sensor. Notably, however, RNAi of the conserved E3 SUMO ligase gene gei-17 (PIAS1/Su(var)2–10) (Hari et al., 2001; Mohr and Boswell, 1999; Ninova et al., 2020) did not desilence the piRNA sensor (Figure 1C).

Null alleles of many of these genes cause embryonic arrest, which precludes an analysis of silencing in the adult germline. To further explore the role of SUMO and HDAC factors in piRNA silencing, we therefore tested whether partial or conditional loss-of-function alleles activate the piRNA sensor. Auxin-inducible degron alleles of hda-1, let-418, mep-1, and mrg-1 and a truncation allele of sin-3 all desilenced the piRNA sensor in 100% of worms examined (Figure 1C). We found that a 3xflag fusion to the endogenous smo-1 gene desilenced the piRNA sensor in ~40% of adults analyzed (Figure 1C), suggesting that this tagged smo-1 allele behaves like a partial loss of function. A strain expressing a temperature-sensitive UBC-9(G56R) protein desilenced the sensor in 90% of the animals at the semi-permissive temperature of 23°C (Figure 1C) (Kim et al., 2021). By contrast, presumptive null alleles of gei-17 completely failed to desilence the piRNA sensor strain (Kim et al., 2021). Together, these findings suggest that histone deacetylase complexes and components of the SUMO pathway promote piRNA-mediated silencing.

SUMOylation of HDA-1 promotes piRNA surveillance

In a parallel study, we found that C. elegans HDA-1 is SUMOylated in the adult germline (Kim et al., 2021). Moreover, we showed that SUMOylation of HDA-1, formation of an adult NuRD complex, and piRNA-mediated silencing depend redundantly on PIE-1, a CCCH zinc finger protein with SUMO E3-like function, and on GEI-17, a homolog of PIAS1/Su(var)2–10 SUMO E3 ligase, suggesting that SUMOylation of HDA-1 might promote piRNA surveillance (Kim et al., 2021). Human HDAC1 is SUMOylated near its C-terminus on consensus SUMO-acceptor lysines 444 and 476 (Figure 2ADavid et al., 2002), but HDA-1 does not contain consensus SUMO acceptor sites, and poor conservation between the C-termini of human HDAC1 and worm HDA-1 did not suggest the identity of potential SUMO-acceptor sites. However, GPS-SUMO prediction software identified lysines 444 and 459 of HDA-1 as possible non-consensus SUMO-acceptor sites (Figure 2AZhao et al., 2014). To investigate whether one or both of these lysine residues is required for HDA-1 SUMOylation, we used CRISPR genome editing to mutate the endogenous hda-1 gene in a strain that expresses SUMO fused to 10 N-terminal histidines, his10::smo-1 (Kim et al., 2021). We then used nickel-nitrilotriacetic acid (Ni-NTA) affinity chromatography under stringent denaturing conditions (Tatham et al., 2009) to capture SUMOylated proteins from worm lysates. The Ni-column eluates were then analyzed by western blotting for HDA-1. SUMOylated HDA-1 was recovered by Ni-NTA affinity chromatography from wild-type lysate and from lysates of each single-site lysine-to-arginine mutant (Figure 2B, lanes 7–9), but was absent when both K444 and K459 were mutated together, HDA-1(KKRR) (Figure 2B, lane 10). As a control, the protein MRG-1, which is highly SUMOylated in wild-type worms (Kim et al., 2021), was readily detected in worms expressing HDA-1(KKRR) (Figure 2B). When we introduced the piRNA sensor into the HDA-1 SUMO-acceptor site mutants, we found that whereas the piRNA sensor remained silent in the single-site mutants (n = 30) (data not shown), it was expressed in 100% of HDA-1(KKRR) worms (Figure 2C).

SUMOylation of HDA-1 at K444 and K459 facilitates piRNA-mediated silencing.

(A) Domain structure of C. elegans type 1 histone deacetylase HDA-1 and C-terminal location of SUMO-acceptor sites. Sequence alignment showing poor conservation at the C-termini of C. elegans HDA-1 and Homo sapiens HDAC1. The human HDAC1 C-terminus possesses two consensus SUMO-acceptor sites, K444 and K476 (acceptor lysines in red; consensus SUMO acceptor motif in pink box). GPS-SUMO predicts two candidate non-consensus SUMOylation sites in HDA-1, both near the C-terminus, K444 and K459 (red lysines in green boxes). (B) Western blot analyses of HDA-1 and MRG-1 before (lanes 1–5) and after (lanes 6–10) affinity enrichment of SUMOylated proteins from wild-type (wt) or hda-1 SUMO acceptor-site mutants. SUMOylated proteins were enriched from worms expressing HIS10::SMO-1. Black arrowheads indicate SUMOylated HDA-1 and MRG-1; white arrowheads indicate unmodified forms of HDA-1 and MRG-1. Asterisks indicate non-specific bands. Additional higher forms (indicated by white star) were detected, suggesting Multi-monoSUMOylation or PolySUMOylation of HDA-1. (C) Analysis of piRNA-mediated silencing in SUMOylation-defective mutants and rescue by HDA-1::SMO-1 translational fusion. (Top) The color of each bar indicates the percentage of worms in which the piRNA sensor was silent (OFF, gray) or expressed (ON, green). Thirty (n = 30) worms of each genotype were examined. (Bottom) Differential interference contrast and epifluorescence images of dissected gonads from hda-1[KKRR] and hda-1[KKRR]::smo-1. (D) Western blots showing levels of HDA-1 and variants proteins expressed from the endogenous hda-1 locus. Tubulin was used as a loading control. (E) Brood size analysis of wt worms or HDA-1 SUMOylation-site mutants, and rescue by HDA-1::SMO-1 translational fusion. Worms were grown at 20°C or 25°C, as indicated. Statistical significance was determined by ordinary one-way ANOVA: *p<0.05; **p<0.01; ****p<0.0001; ns: not significant. (F) Mortal germline analyses of wt or HDA-1 SUMOylation-site mutant, and HDA-1::SMO-1 fusion worms. The wago-9/hrde-1 mutant has a severe mortal germline phenotype. Worms were passaged at 25°C for eight generations, and the average number of progeny from 10 individuals (n = 10) was determined at each generation. Error bars represent standard error of the mean (SEM).

Since the SUMOylation sites in HDA-1 are very close to the C-terminus, we wondered if appending SUMO via translation fusion to the HDA-1(KKRR) mutant protein might restore HDA-1 function in the piRNA sensor assay. Using CRISPR, we inserted a modified smo-1 open reading frame just before the stop codon of the hda-1[KKRR] gene at the endogenous hda-1 locus. The modified SMO-1 fusion cannot be transferred to other proteins because it lacks the GG amino acids required for conjugation (Dorval and Fraser, 2006; see Materials and methods). Surprisingly, the resulting hda-1[KKRR]::smo-1 strain was homozygous viable, healthy, and expressed an HDA-1::SMO-1 fusion protein at levels similar to those observed for wild-type HDA-1 (Figure 2D). Strikingly, appending SMO-1 to the C-terminus of HDA-1(KKRR) completely rescued the silencing defect of HDA-1(KKRR) (Figure 2C). Moreover, HDA-1(KKRR)::SMO-1 rescued the piRNA-mediated silencing defects of smo-1 and ubc-9 mutants (Figure 2C), suggesting that the SUMO pathway promotes piRNA-mediated silencing via C-terminal SUMOylation of HDA-1.

HDA-1 SUMOylation is not required for maintenance of piRNA-initiated silencing

Once silencing is established by the upstream components of the Piwi pathway, it can be maintained indefinitely by the co-transcriptional arm of the pathway without the continued need for prg-1 activity (Shirayama et al., 2012). For example, the initial silencing of the gfp::cdk-1 transgene requires PRG-1 activity, but maintenance of silencing does not (Shirayama et al., 2012). To ask if HDA-1 SUMOylation is required for the maintenance of piRNA-initiated silencing, we crossed an already silenced gfp::cdk-1 transgene into an hda-1[KKRR] mutant strain. We found that the gfp-1::cdk-1 reporter remained silent even after five generations in HDA-1(KKRR) worms (n = 26), supporting the idea that C-terminal SUMOylation of HDA-1 is not required to maintain piRNA-induced transcriptional silencing.

HDA-1 SUMOylation mutants cause temperature-dependent reductions in fertility

Factors that promote genome integrity and epigenetic inheritance are required for germline immortality: their loss causes a mortal germline phenotype, whereby fertility declines in each generation and this decline is often exacerbated at elevated temperatures (Ahmed and Hodgkin, 2000). Although hda-1 is an essential gene required for embryonic development (Shi and Mello, 1998), worms expressing HDA-1(KKRR) and HDA-1(KKRR)::SMO-1 from the endogenous hda-1 locus were viable and fertile. Careful examination of brood size revealed that HDA-1(KKRR) worms made significantly fewer progeny than wild-type worms at 20°C and at 25°C (Figure 2E). When maintained at 25°C, the fertility of HDA-1(KKRR) worms steadily declined over several generations, from an average of 132 progeny in the first generation to fewer than 10 progeny in the fifth and subsequent generations (Figure 2F). Wild-type worms also showed an initial decline in fertility when maintained at 25°C, but averaged ~100 progeny in the fourth and subsequent generations (Figure 2F). By contrast, wago-9/hrde-1 mutants showed a rapid decline in fertility and could not be maintained beyond the third generation (Figure 2FBuckley et al., 2012; Spracklin et al., 2017). Appending SUMO via translational fusion, HDA-1(KKRR)::SMO-1, rescued the fertility defect of HDA-1(KKRR) worms, suggesting that HDA-1 SUMOylation promotes fertility (Figure 2E). When maintained at 25°C, however, HDA-1(KKRR)::SMO-1 worms gradually became infertile over five generations (Figure 2F). Worms expressing HDA-1::SMO-1—with intact SUMO-acceptor sites—were similar to HDA-1(KKRR)::SMO-1 animals, exhibiting significantly reduced fertility at 20°C and a further progressive decline in fertility over five generations at 25°C (Figure 2F). Thus, properly regulated SUMOylation of HDA-1 is essential for germline immortality.

SUMOylation promotes HDA-1 association with other chromatin factors including NuRD complex components

SUMOylation modulates protein interactions (Hendriks and Vertegaal, 2016; Kerscher, 2007). To examine how SUMOylation affects HDA-1 complexes, we introduced a GFP tag into the C-terminus of the endogenous wild-type and mutant hda-1 alleles and used GFP-binding protein (GBP) beads to immunoprecipitate the HDA-1::GFP fusion proteins (Rothbauer et al., 2008). SDS-PAGE analysis revealed that a core set of proteins strongly interact with HDA-1::GFP (Figure 3A). Mass spectrometry (MS) of the corresponding gel slices identified these proteins as: LIN-40, a homolog of metastasis-associated protein (MTA1) (Chen and Han, 2001); LIN-53, a homolog of retinoblastoma-associated protein 46/48 (RBAP46/48) (Solari and Ahringer, 2000); DCP-66, a homolog of GATA zinc finger domain containing protein GATAD (Käser-Pébernard et al., 2014); and SPR-5, a homolog of lysine demethylase (LSD1/KDM1) (Katz et al., 2009).

SUMOylation of HDA-1 promotes its association with NuRD and other chromatin factors.

(A) Silver stained gel of proteins that co-immunoprecipitate with HDA-1::GFP. The indicated protein bands were excised from the gel and identified by mass spectrometry. (B) Scatter plot comparing the levels of proteins identified by mass spectrometry in HDA-1::GFP IPs from smo-1(RNAi) and wild-type (no RNAi) worms. The x axis shows the log value of spectral counts for each protein identified by IP-MS from wild-type worms. The y axis shows the log ratio of spectral counts for each protein in HDA-1::GFP IPs from smo-1(RNAi) vs. wild-type. (C) As in B, but comparing hda-1[KKRR]::gfp to hda-1[WT]::gfp. In (B) and (C), the spectral counts of HDA-1 (RED) were used to normalize between samples. A full list of the identified proteins is provided in Supplementary file 2. (D) Western blot analyses of proteins (indicated to left of blots) that associate with MEP-1::GTF (GFP IP) in hda-1[WT], hda-1[KKRR] and hda-1[KKRR]::smo-1 lysates. The detected proteins are indicated to the right (black arrowheads). The modified isoforms, HDA-1::SUMO and HDA-1::SUMO-UBIQUITIN are indicated with white arrow and white star, respectively. (E) Side-by-side comparison of HDA-1 isoforms detected in the HDA-1, SMO-1, and UBIQUITIN blots in (D). The black dot indicates an unknown HDA-1 isoform. The black star indicates an unknown SUMOylated protein.

We also used reversed-phase high-performance liquid chromatography (RP-HPLC) MS to identify HDA-1::GFP interactors (see Materials and methods). Among approximately 200 high-confidence interactors with ≥10 spectral counts, we identified 63 proteins that were depleted by an arbitrary cutoff of 40% in immunoprecipitates from both smo-1(RNAi) and hda-1[KKRR]::gfp lysates (Figure 3B, C and Supplementary file 2). These SUMO-dependent interactors included MEP-1 (Unhavaithaya et al., 2002), AMA-1 (major subunit of pol lI) (Sanford et al., 1983), and MET-2 (SETDB1 H3K9 histone methyltransferase) (Andersen and Horvitz, 2007; Bessler et al., 2010). This analysis also revealed that the core interactors identified above, LIN-40, LIN-53, DCP-66, and SPR-5, were reduced (by 12–37%) in immunoprecipitations (IPs) from smo-1(RNAi) or HDA-1(KKRR) lysates (Figure 3B, C and Supplementary file 2). Of note, interactions between HDA-1 and SIN-3/SIN3A were not sensitive to SUMO-pathway perturbations (Figure 3B, C and Supplementary file 2).

HDA-1 SUMOylation promotes the association of MEP-1 with chromatin regulators

To explore the molecular consequences of HDA-1 SUMOylation on its physical interactions with MEP-1 and other chromatin regulators, we crossed hda-1[KKRR] and hda-1[KKRR]::smo-1 strains to worms that express a tandemly tagged MEP-1::GFP::TEV::3XFLAG (MEP-1::GTF) from the endogenous mep-1 locus (Kim et al., 2021). We then used GBP beads to immunoprecipitate MEP-1::GTF protein complexes from hda-1 wild-type or mutant lysates. As expected, interactions between MEP-1::GTF and LET-418/Mi-2 did not depend on HDA-1 SUMOylation (Figure 3D; Kim et al., 2021). Consistent with our HDA-1 proteomics studies, however, MEP-1::GTF pulled down wild-type HDA-1 but not HDA-1(KKRR) (Figure 3D). Moreover, LIN-53, MRG-1, and the HP-like heterochromatin proteins HPL-1 and HPL-2 were also greatly reduced in MEP-1 complexes purified from hda-1[KKRR] lysates (Figure 3D). Strikingly, each of these factors were dramatically increased, above wild-type levels, in MEP-1 complexes purified from hda-1[KKRR]::smo-1 lysates (Figure 3D). Thus, the C-terminal fusion of SUMO to HDA-1 rescues MEP-1 interactions with HDA-1(KKRR) and also promotes MEP-1 interaction with HDA-1-binding partners. It is important to note that SUMO-conjugation (in contrast to the SUMO-translational fusion) is rapidly reversed in lysates prepared for IP studies (Kim et al., 2021), which explains why the bands detected by western blotting in Figure 3D—even for the strongly SUMOylated MRG-1—migrate at the size of the unmodified proteins.

In MEP-1 immunoprecipitates from hda-1[KKRR]::smo-1 lysates, we detected multiple HDA-1 bands, including a prominent band slightly larger than the expected size of HDA-1::SMO-1 (white stars in Figure 3D, E). Western blots with ubiquitin-specific antibody suggested that this prominent band is a mono-ubiquitinated form of the fusion protein (Figure 3D, E). In both input and IP samples, we also observed an isoform similar in size to endogenous HDA-1 that was not detected by SUMO-specific antibodies (Figure 3D, E), suggesting that the HDA-1::SMO-1 fusion protein may be cleaved near the C-terminus of HDA-1, removing the SUMO peptide.

HDA-1 SUMOylation promotes histone deacetylation in vivo

The findings above suggest that SUMOylation promotes the assembly and function of HDA-1 complexes. Our proteomic studies also revealed that SUMOylation promotes interactions between HDA-1 and other histone-modifying enzymes required for heterochromatin formation, including the demethylase SPR-5/LSD1, which removes activating H3K4me2/3 marks, and the methyltransferase MET-2/SetDB1, which installs silencing H3K9me2/3 marks (Greer and Shi, 2012). Consistent with the idea that SUMOylation of HDA-1 promotes silencing via SPR-5 and MET-2, immunostaining revealed greatly reduced levels of H3K9me2 and increased levels of the H3K4me3 and throughout the germline in HDA-1(KKRR) worms as compared to wild-type (Figure 4A, B, Figure 4—figure supplement 1). Immunostaining also revealed higher levels of acetylated H3K9 (H3K9Ac) in germlines of HDA-1(KKRR) and mep-1-depleted worms than in wild-type (Figure 4C). Moreover, we found that HDA-1, LET-418, and MEP-1 (including MEP-1 expressed from the germline-specific wago-1 promoter) bind heterochromatic regions of the genome, depleted of the activating H3K9Ac mark and enriched for the silencing marks H3K9me2/3 (Figure 4D). Thus, SUMOylation of HDA-1 appears to drive formation or maintenance of germline heterochromatin.

Figure 4 with 1 supplement see all
HDA-1 SUMOylation is required for formation of germline heterochromatin.

(A, B) Immunofluorescence micrographs of (A) wild-type (wt) and (B) hda-1[KKRR] gonads stained with anti-H3K9me2 antibody and DAPI. The dashed boxes indicated ‘a’ and ‘b’ are enlarged as shown. Two representative gonads are shown for each strain. (C) Differential interference contrast and immunofluorescence micrographs of gonads from wt, hda-1[KKRR], mep-1::gfp::degron with auxin (50 µM) worms stained with anti-H3K9Ac antibody and DAPI. (D) Genome Browser tracks (Integrated Genomics Viewer [IGV]) showing ChIP-seq peaks for NuRD complex components (HDA-1, LET-418, and MEP-1) and three histone modifications (H3K9Ac, H3K9me2, and H3K9me3) along each C. elegans chromosome (I–V and X). MEP-1(gonad) data are from worms that express MEP-1::GTF only in the germline, using the wago-1 promoter, for germline-specific CHIP.

SUMOylated HDA-1 and PRG-1 co-regulate hundreds of targets, including many spermatogenesis genes

The visibly reduced level of germline heterochromatin in HDA-1(KKRR) worms suggests that gene expression is broadly misregulated when HDA-1 cannot be SUMOylated. To examine the effect of HDA-1 SUMOylation on germline gene expression, we performed high-throughput sequencing of mRNAs isolated from dissected germlines of hda-1[KKRR], hda-1[KKRR]::smo-1, and ubc-9[G56R] animals, and from worms expressing degron alleles of hda-1 and mep-1 with or without auxin exposure beginning at the L4 stage (Figure 5A). Replicate libraries gave highly reproducible mRNA profiles from each mutant (Figure 5—figure supplement 1A). Depletion of HDA-1 or MEP-1 or inactivation of the SUMO pathway caused widespread upregulation of germline mRNAs and transposon families, with extensive but incomplete overlap between the mutants (Figure 5B, Figure 5—figure supplement 1B, Figure 5—figure supplement 2). Twice as many genes were upregulated in degron::hda-1 germline as in degron::mep-1 or ubc-9[G56R] (Figure 5B, Figure 5—figure supplement 1B), likely reflecting the role of HDA-1 in multiple complexes. Nearly 10-fold fewer genes were upregulated in hda-1[KKRR] than in auxin-treated degron::hda-1 animals (Figure 5B, Figure 5—figure supplement 1B), consistent with the phenotypic differences between the two mutants. Most (305, ~71%) of the genes upregulated in hda-1[KKRR] germlines were also upregulated in auxin-treated degron::hda-1 animals (Figure 5B). As expected, mRNAs upregulated in hda-1[KKRR] were restored to nearly wild-type levels in hda-1[KKRR]::smo-1 (Figure 5C, D). Consistent with the known roles of the NuRD complex and SUMOylation pathways in modulating chromatin states, we observed a loss of enrichment for H3K9me2, as measured by ChIP-seq, near the promoters of genes upregulated in hda-1[KKRR], ubc-9[G56R], and auxin-treated degron::hda-1 worms (Figure 5—figure supplement 3).

Figure 5 with 5 supplements see all
The SUMO, NuRD, and piRNA pathways regulate the same group of targets.

(A) Schematic of mRNA-seq from dissected gonads. (B) Venn diagram showing overlap between upregulated genes in degron::hda-1, mep-1::degron, ubc-9(G56R), and hda-1[KKRR] germlines. Numbers in parentheses indicate total number of upregulated genes. (C) Scatter plot of upregulated genes in hda-1[KKRR]. The x-axis represents reads in wild-type (wt), and y-axis represents reads in hda-1[KKRR]. (D) Scatter plot showing the effect of hda-1[KKRR]::smo-1 on the 430 genes (from C) upregulated in hda-1[KKRR]. The x-axis represents reads in wt, and y-axis represents reads in hda-1[KKRR]::smo-1. (E) Venn diagram showing overlap between upregulated genes in hda-1[KKRR], prg-1, rde-3, and wago-9 mutants. (F) Bar graph showing fractions of upregulated genes involved in spermatogenesis, oogenesis, neutral, or other categories. ‘Other’ indicates genes that cannot be put into one of the other categories (Ortiz et al., 2014). Genes with >1 mRNA-seq reads in wt gonad were used to generate the ‘wild-type’ dataset as a reference. The number of genes in each dataset is labeled at the top. (G–I) Scatter plots comparing mRNA-seq reads in (G) hda-1[KKRR], (H) rde-3, and (I) prg-1(ne4766) to those in wt. The blue dashed lines indicate a twofold increase or decrease in mutant compared to wt.

Because HDA-1 SUMOylation is required for silencing of a piRNA reporter, we examined which endogenous mRNAs are co-regulated by HDA-1 SUMOylation and the piRNA pathway factors, PRG-1, WAGO-9/HRDE-1, and RDE-3. We prepared mRNA sequencing libraries from dissected gonads collected from prg-1(ne4766), wago-9/hrde-1(tm1200), and rde-3(ne3370) mutant animals (Figure 5A). RDE-3 is required for production of small RNAs (termed 22G-RNAs) that guide transcriptional and post-transcriptional silencing by WAGO Argonautes, including WAGO-9 (Gu et al., 2009; Zhang et al., 2011). Of the 430 mRNAs upregulated (≥2-fold) in hda-1[KKRR], we found that 362 (84%) were upregulated (≥2-fold) in prg-1(ne4766), 360 (84%) were upregulated in rde-3(ne3370), and 331 (77%) were upregulated in all three mutant strains (Figure 5E). Similarly, the genes upregulated in hda-1[KKRR] accounted for 40% of the genes upregulated in prg-1(ne4766) and 50% of those upregulated in rde-3(ne3370) (Figure 5E). Fewer genes were upregulated in wago-9/hrde-1 (Figure 5E, Figure 5—figure supplement 1B), perhaps due to redundancy with other WAGO Argonautes. Whereas only 5 transposon families were upregulated in prg-1(ne4766) and a total of 6 transposon families were upregulated in hda-1[KKRR] (Figure 5—figure supplement 2), thirty (30) were upregulated in rde-3 mutants (Figure 5—figure supplement 2), consistent with the previously described role for rde-3 and the WAGO pathway in maintaining the silencing of most transposons in worms. The silencing defect was more severe in auxin-treated degron::hda-1 than in hda-1[KKRR] (Figure 5—figure supplement 1B, Figure 5—figure supplement 2), resulting in the increased expression of many more transposons and a more extensive overlap with genes upregulated in rde-3 mutant worms (Figure 5—figure supplement 1C, Figure 5—figure supplement 2). This result indicates that HDA-1 also promotes the maintenance of silencing independently of HDA-1 SUMOylation.

Most of the genes upregulated in prg-1, rde-3, and hda-1[KKRR] mutants are normally expressed during spermatogenesis (Figure 5F–I, Figure 5—figure supplement 4). In most cases, the upregulation of these spermatogenesis mRNAs did not correlate with reduced WAGO 22G-RNAs targeting these genes (Figure 5—figure supplement 5), suggesting that the spermatogenesis switch may be regulated indirectly by the small RNA pathways.

HDA-1 physically interacts with WAGO-9/HRDE-1 and functions in inherited RNAi

Because WAGO-9/HRDE-1 is a nuclear Argonaute that functions downstream in the Piwi pathway to establish and maintain epigenetic silencing (Ashe et al., 2012; Bagijn et al., 2012; Buckley et al., 2012; Shirayama et al., 2012), we asked if WAGO-9/HRDE-1 interacts with HDA-1. We used CRISPR genome editing to generate a functional gfp::wago-9 strain and then used GBP beads to immunoprecipitate GFP::WAGO-9 complexes. Western blot analyses revealed that HDA-1 co-precipitates specifically with GFP::WAGO-9 (Figure 6A). We also found that the HP1-like protein HPL-2 interacts with WAGO-9/HRDE-1 (Figure 6A), consistent with previous genetic studies (Ashe et al., 2012; Buckley et al., 2012; Gu et al., 2012; Luteijn et al., 2012; Shirayama et al., 2012). HPL-2 binds methylated H3K9 in heterochromatin (Garrigues et al., 2015). Neither HDA-1 nor HPL-2 were precipitated by GBP beads incubated with lysates prepared from untagged wild-type worms. These interactions were confirmed by reciprocal GFP IP experiments using hda-1::gfp lysates (Figure 6B). Moreover, the interaction of HDA-1 with WAGO-9 and HPL-2 was reduced in hda-1[KKRR] animals or by smo-1(RNAi) (Figure 6B). Interestingly, we observed a truncated form of HDA-1(KKRR)::GFP in the input samples that appears to result from proteolytic removal of the C-terminal GFP (Figure 6B, filled arrow, compare input to IP lanes). By contrast, GFP is not proteolytically removed from wild-type HDA-1::GFP in the presence or absence of smo-1(RNAi). The reason for this difference in protein stability is not known and will require further study. Taken together, the SUMO-dependent interactions between HDA-1 and WAGO-9 suggest that SUMOylation of HDA-1 promotes the assembly of an Argonaute-guided nucleosome-remodeling complex.

HDA-1 interacts with WAGO-9/HRDE-1, which requires HDA-1 SUMOylation.

(A) Western blot analysis showing that HDA-1 and HPL-2 co-immunoprecipitate with GFP::WAGO-9/HRDE-1. (B) Western blot analysis of WAGO-9 and HPL-2 in HDA-1::GFP immunoprecipitates from wild-type, HDA-1 SUMO acceptor-site mutant, or smo-1(RNAi) worms. Blotting with anti-SMO-1 antibody showed depletion of SMO-1 in the smo-1(RNAi) worms. A band the size of untagged HDA-1 (black arrowhead) in hda-1[KKRR]::gfp in input appears to be a cleavage product that removes the GFP tag. (C) Graph showing the levels of silencing induced by RNAi over three generations in wild-type, hda-1[KKRR], and hda-1[KKRR]::smo-1 worms. Worms were treated with gfp(RNAi) at P0, and F1 larvae were transferred to regular NGM plates. The percentage of worms that express OMA-1::GFP was scored (n≥22, two replicates). Error bars represent standard error of the mean (SEM).

Many of the factors that maintain heritable epigenetic silencing triggered by piRNAs—including WAGO-9/HRDE-1—are also required for multigenerational germline silencing triggered by exogenous dsRNA, that is, inherited RNAi (Ashe et al., 2012; Buckley et al., 2012; Shirayama et al., 2012). We therefore asked if HDA-1 SUMOylation is also required for inherited RNAi. Worms expressing a bright germline OMA-1::GFP were fed bacteria that express GFP dsRNA (i.e., RNAi by feeding) for one generation (P0). The subsequent F1 and F2 generations were removed from the RNAi food and cultured on a normal Escherichia coli diet (no gfp dsRNA). In wild-type worms, oma-1::gfp was silenced in the P0 generation (in the presence of gfp dsRNA) and remained silent in the F1 and F2 generations in the complete absence of gfp dsRNA (Figure 6C). By contrast, as previously shown (Ashe et al., 2012; Buckley et al., 2012; Spracklin et al., 2017), oma-1::gfp is efficiently silenced in P0 wago-9/hrde-1 worms in the presence of gfp dsRNA, but oma-1::gfp is reactivated in the F1 and F2 generation after the removal from gfp dsRNA. hda-1[KKRR] animals were similarly defective for heritable RNAi: oma-1::gfp was efficiently silenced in the P0 worms in the presence of gfp dsRNA, but expression was fully restored in the F1 and F2 generations in the absence of gfp dsRNA (Figure 6C). Finally, the hda-1[KKRR]::smo-1 translational fusion fully rescued the heritable RNAi defect of hda-1[KKRR] (Figure 6C). Taken together, these findings suggest that HDA-1 SUMOylation promotes heritable transcriptional silencing in both the piRNA and RNAi pathways.

Discussion

HDAC1 SUMOylation promotes Argonaute-directed transcriptional silencing

Argonaute small RNA pathways collaborate with chromatin factors to co-regulate gene expression, transposon activity, and chromosome dynamics (reviewed in Almeida et al., 2019). Here, we have identified components of the NuRD complex as well as the SUMO and its E2 protein ligase, UBC-9, as chromatin-remodeling and -modifying complexes required for the initiation of Piwi-mediated gene silencing. Using mutagenesis of candidate SUMO-acceptor sites and affinity enrichment of SUMO, we identified C-terminal lysines required for the SUMOylation of the HDAC1 homolog, HDA-1. Mutating these lysines to arginine in HDA-1(KKRR) abolished the initiation but not the maintenance of piRNA-mediated silencing and also abolished the initiation of transgenerational silencing in response to dsRNA. Appending SUMO by translational fusion to HDA-1(KKRR) completely rescued Argonaute-mediated silencing and even restored piRNA silencing in animals with mutations in the SUMO conjugating machinery. Taken together, these molecular genetic studies strongly implicate HDAC1 SUMOylation in promoting Argonaute-directed transcriptional silencing (Figure 7).

Model: HDAC1 SUMOylation promotes Argonaute-directed transcriptional gene silencing.

SUMOylation of HDA-1 enables nucleosome remodeling and deacetylase (NuRD) complex assembly in the adult germline and WAGO-9 or other Argonautes recruit the NuRD complex to piRNA targets. The Argonaute/NuRD complex, along with other histone-modifying enzymes—for example, SPR-5, MET-2—removes the active histone marks (H3K9Ac and H3K4me2/3) and establishes silencing marks (H3K9me2/3) to suppress their transcription.

SUMO: a potent genetic modifier with an elusive biochemical signature

Given the strong genetic evidence that the modification of HDA-1 by SUMO promotes piRNA silencing in the adult germline, we were surprised that the conjugated isoform of HDA-1 was undetectable in our IP assays from adult animals. Only when Ni-Affinity purification was used under strict denaturing conditions was it possible to recover the HDA-1-SUMO isoform. A likely explanation for these findings is that the covalent attachment of SUMO to its target proteins is rapidly reversed in lysates from C. elegans adults (Kim et al., 2021).

The dynamic nature of SUMO conjugation is well known from studies on other organisms (Mukhopadhyay and Dasso, 2007), and the SUMO protease SENP1 has been identified as an important regulator of human HDAC1 SUMOylation (Cheng et al., 2004, see more discussion below). In the future, it will be interesting to explore whether any of the three C. elegans homologs of SENP1 regulate HDA-1 SUMOylation in the worm germline. The presence of such factors, which appear to be very active in worm lysates, likely explains why so little HDA-1-SUMO was recovered in our IP studies. Indeed, it seems likely that the labile nature of the SUMO interaction could explain the discordance between its robust genetic effect and its surprisingly weak physical association with its targets in co-IP assays. Consistent with this idea, an HDA-1::SMO-1 translational fusion protein with an abnormal linkage via the N-terminus of SUMO was stable in protein lysates, strongly rescued the silencing defects of the presumptive SUMO-acceptor mutant protein HDA-1(KKRR), and dramatically enhanced the detection of protein-protein interactions between HDA-1 and components of an adult-stage NuRD complex.

We were surprised that the HDA-1::SMO-1 fusion protein, which was constructed at the endogenous and essential hda-1 locus, was so well-tolerated. Conceivably, the cleavage and ubiquitination of HDA-1::SMO-1 that we detected in our IP assays provide alternative mechanisms to counter the effects of this translational fusion. The mono-ubiquitinated form of the HDA-1::SMO-1 protein exhibited reduced staining with a SUMO-specific antibody, raising the possibility that the SUMO moiety in this protein is itself modified by ubiquitin, a modification that has been reported for SUMO isoforms in humans (Danielsen et al., 2011; Tatham et al., 2008). Taken together, our findings suggest that HDA-1 SUMOylation is not only important genetically but is highly dynamic and may be regulated at multiple levels.

HDAC1 SUMOylation plays diverse roles in gene regulation

Studies in mice and in human cell culture have investigated SUMOylation of C-terminal HDAC1 lysines (Cheng et al., 2004; Citro et al., 2013; David et al., 2002; Joung et al., 2018; Tao et al., 2017). In at least three cases, mutating these lysines increased the levels of histone acetylation and transcriptional activation (David et al., 2002; Cheng et al., 2004; Joung et al., 2018). As mentioned above, the SUMO protease SENP1 was identified as a factor that promotes the removal of C-terminal SUMO moieties from human HDAC1 in a prostate cancer model, causing upregulation of the androgen receptor (Cheng et al., 2004). In a subsequent study, these authors identified SENP1 as upregulated in human prostate cancers and found that overexpression of SENP1 was sufficient to drive prostate neoplasia in a mouse model (Cheng et al., 2006). One very interesting study showed that Aβ insult led to upregulation of the SUMO E3 factor PIAS and resulted in SUMOylation of C-terminal lysines in HDAC1 in the rat hippocampus (Tao et al., 2017). Remarkably, this study showed that a lentivirus-driven HDAC1::SUMO translational fusion protein could rescue the learning and memory deficits of an APP/Presenilin 1 murine model of Alzheimer's disease (Tao et al., 2017), suggesting that HDAC1 SUMOylation may be neuroprotective in response to Aβ accumulation. Taken together, our worm studies and these studies in mammalian systems point to the likely importance of HDAC1 SUMOylation in the regulation of gene expression in diverse animals.

SUMO promotes the assembly of a germline MEP-1-NuRD complex

Precisely how HDA-1 SUMOylation promotes its downstream functions in piRNA silencing will require further work. One possibility is that SUMO acts as a bridge between HDA-1 and a nuclear Argonaute WAGO-9, while simultaneously promoting NuRD complex assembly (Figure 7 model). This model is supported by the dramatically enhanced association between MEP-1 and the HDA-1::SMO-1 translational fusion and by the ability of HDA-1::SMO-1 to rescue the piRNA silencing defects of upstream SUMO pathway mutants.

Paradoxically, HDA-1 is not SUMOylated in the embryo and yet interacts robustly with MEP-1 (Kim et al., 2021). We do not know why the association of HDA-1 with MEP-1 requires SUMO in the adult germline but not the embryo. Perhaps unknown co-factors promote their SUMO-independent interaction in the embryo. Alternatively, adult germline-specific factors may inhibit or modify HDA-1 or other NuRD complex components to prevent their direct association. SUMOylation of HDA-1 in the adult may therefore enable it to associate with MEP-1 via a mechanism bridged by SUMO, a SUMO glue, or SUMO-SIM network mode of interaction (Matunis et al., 2006; Psakhye and Jentsch, 2012 Pelisch et al., 2017). The MEP-1 protein has two consensus SIMs, which could be important for this association between MEP-1 and HDA-1. However, our preliminary studies on these motifs were confounded by the finding that their simultaneous mutation resulted in a completely sterile inactive mep-1 allele, similar phenotypically to a null allele. Thus, although the SIM motifs in MEP-1 may be important for its interaction with HDA-1-SUMO, they may also be required for other functions or for interactions with other essential co-factors.

Parallels in the role of SUMOylation in Piwi silencing in insects, mice, and worms

Histone deacetylation is a necessary step in de novo transcriptional silencing. Yet, precisely how nuclear Argonautes orchestrate both deacetylation and the subsequent installation of silencing marks on target chromatin is not known. The nuclear Argonaute WAGO-9/HRDE-1 initiates transcriptional silencing downstream of both the piRNA- and the dsRNA-induced silencing pathways (Ashe et al., 2012; Bagijn et al., 2012; Buckley et al., 2012; Shirayama et al., 2012), and we have shown here that HDA-1 interacts with WAGO-9 in a manner that partially depends on HDA-1 SUMOylation. In fruit fly, the nuclear Argonaute Piwi was recently shown to interact with the SUMO E3 PIAS homolog Su(var)2–10 and to promote the recruitment of the histone methyltransferase SetDB1/EGGless (Egg) and its cofactor MCAF1/Windei (Wde) (Ninova et al., 2020). A worm paralog of EGG, MET-2, was identified in our SUMO-dependent HDA-1 IP complexes. Although the worm PIAS/Su(var)2–10 homolog, GEI-17, scored negative in our piRNA sensor screen, in a parallel study we have identified a synthetic piRNA silencing defect between gei-17 and mutations in pie-1, which encodes a tandem zinc finger protein with properties consistent with SUMO E3 activity (Kim et al., 2021). Thus, an attractive possibility is that upon binding to nascent target transcripts, nuclear Argonautes recruit SUMO E3 factors to promote the SUMOylation and recruitment of both histone deacetylase and histone methyltransferase complexes.

Interestingly, in Drosophila Kc167 cells, a MEP-1:Mi-2 complex (called MEC) reportedly forms without HDAC1 (Kunert et al., 2009). Kc167 cells are derived from ovarian somatic cells and express components of the Piwi pathway (Vrettos et al., 2017), raising the possibility that HDAC1 SUMOylation might promote its association with the MEC complex in these cells. Interestingly, a recent paper showed that Drosophila MEP-1 and the HDA-1 homolog RPD3 interact with PIWI and with Su(var)2–10 in fly ovarian somatic cells, where they function together to promote transposon silencing (Mugat et al., 2020). Another recent study found that the mouse Piwi homolog MIWI engages NuRD complex components and DNA methyltransferases to establish de novo silencing of transposons in the mouse testes, suggesting additional parallels to the findings from worms and flies (Zoch et al., 2020). It will be interesting in the future to learn if mammalian and insect Piwi Argonautes target HDAC1 SUMOylation to promote de novo Piwi silencing.

The mep-1 gene was previously implicated in regulating the transition from spermatogenesis to oogenesis in hermaphrodites (Belfiore et al., 2002), and our findings suggest that HDA-1 SUMOylation also promotes this function. We were surprised, however, to find that spermatogenesis targets are also regulated by the WAGO-pathway factor RDE-3 and by the Piwi Argonaute PRG-1. Similar findings were reported recently by Reed et al., 2020. These observations raise the interesting possibility that C. elegans germline, Argonaute systems function with SUMO and HDAC1 in promoting the switch from spermatogenesis to oogenesis in hermaphrodites.

Materials and methods

C. elegans strains and genetics

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All the strains in this study were derived from Bristol N2 and cultured on nematode growth media (NGM) plates with OP50, and genetic analyses were performed essentially as described (Brenner, 1974). The strains used in this study are listed in Supplementary file 3.

RNAi screen

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RNAi screen was performed against all 337 genes in the chromatin subset of the C. elegans RNAi collection (Ahringer). RNAi of smo-1 and ubc-9 were added to the screen as chromatin regulators. Synchronous L1 worms of the reporter strain were plated on the isopropyl-β-D-thiogalactoside (IPTG) plates with the corresponding RNAi food. Bacteria with empty L4440 vector served as negative control. The desilencing phenotype was scored when the worms on the control plates grew to young adult stage at 20°C. In the first round of the screen, 78 RNAi clones scored positive. In the second round, we required that the sensor be desilenced in more than 5% of worms, resulting in 29 positive clones, including smo-1 and ubc-9 (Supplementary file 1).

CRISPR/Cas9 genome editing

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The Co-CRISPR strategy (Kim et al., 2014) and mep-1 sgRNA and smo-1 sgRNA (see Kim et al., 2021) were used to generate mep-1::gfp::degron and 3xflag::smo-1 strains. Other CRISPR lines were generated by Cas9 ribonucleoprotein (RNP) editing (Dokshin et al., 2018) or Cas12a (cpf1) RNP editing. Cas12a genome editing mixture containing Cas12a protein (0.5 μl of 10 µg/µl), two crRNAs (each 2.8 μl of 0.2 μg/μl), annealed PCR donor (4 μg), and rol-6(su1006) plasmid (2 μl of 500 ng/μl) was incubated at 37°C for 30 min and 25°C for 1 hr or overnight before injecting animals. For short insertions, like FLAG, auxin-inducible degron, HIS10, and point mutations, synthetic single-strand DNAs were used as donors; for long insertions, like GFP, 2/3xFLAG-Degron, and SUMO, annealed PCR products were used as donors. Guide RNA sequences and ssOligo donors used in this study are listed in Supplementary file 4.

Generation of strains expressing SUMO-conjugated HDA-1

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To prevent transfer of SUMO from the HDA-1::SMO-1 fusion to other SUMO targets, we mutated the tandem C-terminal glycines of SMO-1/SUMO to alanines (GG to AA) (Dorval and Fraser, 2006). The modified smo-1 open reading frame was fused directly before the stop codon of hda-1 or hda-1(ne4747[KKRR]) gene at the endogenous hda-1 locus by CRISPR genome editing as described above.

Auxin treatment

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For the auxin-inducible degron system (Zhang et al., 2015), tir-1::mRuby was expressed in the germline under the control of the sun-1 promoter and eft-3 3′ UTR. The degron-tagged L1 larvae were plated on NGM plates with 100 μM auxin indole-3-acetic acid (IAA; Alfa Aesar, A10556) unless otherwise stated and kept in dark. Worms were collected at young adult stage for further analysis.

Gonad fluorescent image

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Gonads were dissected on glass slide (Thermo Fisher Scientific, 1256820) in M9 buffer, mounted in 2% paraformaldehyde (Electron Microscopy Science, Nm15710) in egg buffer (25 mM HEPES pH 7.5, 118 mM NaCl, 48 mM KCl, 2 mM CaCl2, 2 mM MgCl2), and directly imaged. Epifluorescence and differential interference contrast (DIC) microscopy were performed using an Axio Imager M2 Microscope (Zeiss). Images were captured with an ORCA-ER digital camera (Hamamatsu) and processed using Axiovision software (Zeiss).

Immunofluorescence

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Immunostaining of gonads was performed essentially as described (Kim et al., 2021). Primary antibodies (diluted 1:100) included anti-acetyl-histone H3K9 antibody (Abcam, ab12179), anti-di-methyl-histone H3K9 antibody (Abcam, ab1220), and anti-tri-methyl histone H3K4 (Millipore, 07-473). Secondary antibodies (diluted 1:1000) included goat anti-mouse IgG (H+L) Alexa Fluor 594 (Thermo Fisher Scientific, A11005), goat anti-mouse IgG (H+L) Alexa Fluor 488 (Thermo Fisher Scientific, A11001), and goat anti-rabbit IgG (H+L) Alexa Fluor 568 (Thermo Fisher Scientific, A11011). Epifluorescence and DIC microscopy were performed using an Axio Imager M2 Microscope (Zeiss). Images were captured with an ORCA-ER digital camera (Hamamatsu) and processed using Axiovision software (Zeiss).

Affinity chromatography of histidine-tagged SUMO

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Synchronous adult worms (~200,000) were used for Ni-NTA pull-downs. HIS-tagged SUMO/SUMOylated proteins were enriched as described in Kim et al., 2021.

Co-IP and western blotting

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IPs were performed as described (Kim et al., 2021). For western blots, protein samples were denatured in NuPAGE LDS sample buffer (4×) (Thermo Fisher Scientific, NP0008), loaded on precast NuPAGE Novex 4–12% Bis-Tris protein gel (Life Technologies, NP0321BOX), and transferred onto 0.2 µm nitrocellulose membrane (Bio-Rad, 1704158) using a Transblot turbo transfer system (Bio-Rad). Membranes were incubated with primary antibodies at 4°C overnight and then with secondary antibodies for 1.5 hr at room temperature. Primary antibodies (and dilutions) included anti-FLAG (1:1000) (Sigma-Aldrich, F1804), anti-GFP (1:1000) (GenScript, A01704) anti-MRG-1 (1:1000) (Novus Biologicals, 49130002), anti-HPL-1 (1:1000) (Novus Biologicals, 38620002), anti-HPL-2 (1:1000) (Novus Biologicals, 38630002), anti-LIN-53 (1:1000) (Novus Biologicals, 48710002), anti-HDA-1 (1:2500) (Novus Biologicals, 38660002), anti-LET-418 (1:1000) (Novus Biologicals, 48960002), anti-SMO-1 (1:500) (purified from Hybridoma cell cultures, the Hybridoma cell line was gift from Ronald T. Hay, University of Dundee) (Pelisch et al., 2014), anti-tubulin (1:2000) (Cell Signaling Technology, 9099S), anti-ubiquitin (1:1000) (Abcam, ab7780), and anti-WAGO-9/HRDE-1 (1:500) (gift from Eric A. Miska) (Ashe et al., 2012). Antibody binding was detected with HRP-conjugated secondary antibodies: goat anti-mouse (1:2500) (Thermo Fisher Scientific, 62-6520) and mouse anti-rabbit (1:3000) (Abcam, ab99697).

Immunoprecipitation-mass spectrometry (IP-MS)

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Synchronous adult worms were used for IP experiments unless otherwise indicated. Worm lysates were prepared by combining 1 ml frozen worm pellet mixed with 1 ml IP lysis buffer (Kim et al., 2021) and 2 ml glass beads, and then vortexing on FastPrep (MP Biomedicals) at a speed of 6 m/s for 20 s for four times. Lysates were clarified twice by centrifugation for 30 min each at 12,000 rpm, 4°C. GBP beads were prepared by conjugating GBP nanobody to N-hydroxysuccinimide (NHS)-activated Sepharose beads (Rothbauer et al., 2008). GBP beads were incubated with lysates for 1 hr at 4°C on a rotating shaker, and GFP-tagged proteins and complexes were eluted with 1% sodium dodecyl sulfate (SDS), 50 mM Tris, pH 8.0 at 95°C for 10 min. Proteins were precipitated with trichloroacetic acid (TCA) and digested with trypsin. The resulting peptides were analyzed on a Q Exactive mass spectrometer (Thermo Fisher Scientific) coupled with an Easy-nLC1000 liquid chromatography system (Thermo Fisher Scientific). Peptides were loaded on a pre-column (75 μm ID, 6 cm long, packed with ODS-AQ 120 Å−10 μm beads from YMC Co., Ltd.) and separated on an analytical column (75 μm ID, 13 cm long, packed with Luna C18 1.8 μm 100 Å resin from Welch Materials) using a 78 min acetonitrile gradient from 0% to 30% (v/v) at a flow rate of 200 nl/min. The top 15-most intense precursor ions from each full scan (resolution 70,000) were isolated for HCD MS2 (resolution 17,500; NCE 27), with a dynamic exclusion time of 30 s. We excluded the precursors with unassigned charge states or charge states of 1+, 7+, or >7+ . Database searching was done by pFind 3.1 (http://pfind.ict.ac.cn/) against the C. elegans protein database (UniProt WS235). The filtering criteria were: 1% false discovery rate (FDR) at both the peptide level and the protein level; precursor mass tolerance, 20 ppm; fragment mass tolerance, 20 ppm; the peptide length, 6–100 amino acids. Spectral counts of HDA-1 in IP samples were used for normalization. Proteins either absent in N2 or with more than twofold of the spectra counts in the hda-1::gfp IP compared to those in N2 are shown in Supplementary file 2.

Silver staining

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One-third of each sample from each IP-MS experiment was fractionated on a 4–12% SDS-PAGE gel. Gels were silver stained using ProteoSilver Plus Silver Stain Kit (Sigma-Aldrich). Visible bands were cut for trypsin digestion and MS identification, and the most abundant protein in each band is labeled in Figure 3A.

RNAi inheritance

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L1 oma-1::gfp larvae were placed on IPTG plates (NGM plate with 1 mM IPTG and 100 μg/ml ampicillin) seeded with E. coli strain HT115 transformed with either control vector L4440 or gfp RNAi plasmid. The worms were scored for OMA-1::GFP signal in the oocyte at gravid adult stage and transferred to regular NGM plates. OMA-1::GFP was monitored at gravid adult stage for every generation until most worms recovered the expression of oma-1::gfp.

Germline mortal assay

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For each strain, 18 worms at L4 stage were singled and grown at 25°C. For each generation, the mothers were transferred to new plates every two days and their brood sizes were scored. One worm from each plate for every generation was randomly picked to continue scoring the brood size until plates became totally sterile.

Gonad mRNA-seq and analysis

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For every mutant, about 100 gonads were dissected from the young adult worms (first day as adults). We carefully cut near the −1 oocyte to make sure every gonad is similar and transfer gonads into a 1.5 ml microcentrifuge tube. RNA was extracted from dissected gonads using Tri-reagent with a yield about 0.5 μg. Ribosomal RNA was depleted by RNase H digestion after being annealed with home-made anti-rRNA oligos for C. elegans. DNA was removed by DNase treatment. RNA-seq libraries were constructed using a KAPA RNA HyperPrep kit, and paired-end sequencing was performed on a Nextseq 500 Sequencer with the illumina NextSeq 500/550 high-output kit v2.5 (150 cycles).

Salmon was used to map the mRNA-seq reads with the worm database WS268, and its output files were imported to DESeq2 in R. The differentially expressed genes were filtered by fold change more than 2 and adjusted p-value <0.05. The scatter plots were generated by the plot function in R.

CHIP-seq

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CHIP was carried out using a previous described protocol (Askjaer et al., 2014). Young adult worms were washed three times with M9 and once with PBS and then cross-linked with 1.1% formaldehyde in phosphate-buffered saline (PBS) with protease inhibitors for 10 min before being quenched with 125 mM glycine. Exchanging to CHIP Cell lysis buffer (20 mM Tris-Cl, 85 mM KCl, 0.5% NP40, pH 8.0), DNA was fragmented by sonication (Bioruptor, high intensity, 30 s on and 30 s off for 45 cycles). Samples were incubated with antibodies overnight at 4°C, and then with magnetic beads, which were precleaned with 5% bovine serum albumin (BSA) in PBS + 0.02% Tween-20 (PBST) for 4 hr. After a series of washes in buffers of different stringency, DNA was eluted with CHIP elution buffer (1% SDS, 250 mM NaCl, 10 mM Tris pH 8.0, 1 mM EDTA) at 65°C for 15 min twice. RNA and proteins were removed by RNase A and Proteinase K treatments. Samples were reverse cross-linked by incubating at 65°C, and the DNA was purified using a Zymo DNA clean kit (cat # D5205). Libraries were prepared with NEBNext Ultra II DNA Library Prep Kit. Libraries were multiplexed and sequenced on HiSeq X or NovaSeq 6000 with paired-end 150 bp sequencing. With a pair-wised IP sample and its input, a ChIP-Seq Pipeline in the DolphinNext platform (built by the Bioinformatics core at UMass Medical School) was used to analyze the CHIP-seq data. The data processing and analysis pipeline includes adapter removal (cutadapt), reads mapping (Bowtie2-align-s v2.2.3), duplicates removal (Picard-tools v1.131), peak calling (MACS2 v2.1.1.20160309), peak location, and quantification (Bedtools v2.25.0, Quinlan and Hall, 2010). The output bed files were used to generate figures with IGV (v2.7.2) for Figure 4D. Background subtraction was applied with the intersect function of Bedtools for Figure 5—figure supplement 3.

Small RNA cloning and data analysis

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The small RNA cloning was conducted as the previous report (Shen et al., 2018). Worms were synchronized and collected at young adult stage. Small RNAs were enriched using a mirVana miRNA isolation kit (Invitrogen) from Trizol purified total RNA. Homemade PIR-1 was used to remove the d or triphosphate at the 5′ to generate 5′ monophosphorylated small RNA. Products were then ligated to 3′ adaptor (/5rApp/TGGAATTCTCGGGTGCCAAGG/3ddC/) by T4 RNA ligase 2(NEB) and 5′ adaptor (rGrUrUrCrArGrArGrUrUrCrUrArCrArGrUrCrCrGrArCrGrArUrCrNrNrNrCrGrArNrNrNrUrArCrNrNrN, N is for a random nucleotide) by T4 ligase 1(NEB) sequentially, followed by reverse transcription with RT primer (CCTTGGCACCCGAGAATTCCA) and SuperScript III (Invitrogen). PCR amplification was done with Q5 and primers with indexes (forward: AATGATACGGCGACCACCGAGATCTACACGTTCAGAGTTCTACAGTCCGA, reverse: CAAGCAGAAGACGGCATACGAGAT [6bases index] GTGACTGGAGTTCCTTGGCACCCGAGAATTCCA). PCR productions around 150 bp were separated by 12% SDS-PAGE, extracted with TE buffer (10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0), and purified with isopropanol precipitation. Libraries were equally mixed and sequenced on a NextSeq 550 sequencer using the Illumina NextSeq 500/550 high-output kit v2.5 (75 cycles) with 75 bp single-end sequencing. Adapters were trimmed by cutadapt and reads were mapped to a worm database (WS268) using Bowtie2. DESeq2 was used to normalized reads between samples.

Appendix 1

Appendix 1—key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
AntibodyMouse monoclonal anti-FLAG M2Sigma-AldrichCat# F1804; RRID:AB_262044IB(1:1000)
AntibodyRabbit polyclonal anti-GFPGenScriptCat# A01704; RRID:AB_2622199IB(1:1000)
AntibodyRabbit polyclonal anti-MRG-1Novus BiologicalsCat# 49130002; RRID:AB_10011724IB(1:1000)
AntibodyRabbit polyclonal anti-HPL-1Novus BiologicalsCat# 38620002;
RRID:AB_10008562
IB(1:1000)
AntibodyRabbit polyclonal anti-HPL-2Novus BiologicalsCat# 38630002;
RRID:AB_10647696
IB(1:1000)
AntibodyRabbit polyclonal anti-LIN-53Novus BiologicalsCat# 48710002;
RRID:AB_10011629
IB(1:1000)
AntibodyRabbit polyclonal anti-HDA-1Novus BiologicalsCat# 38660002;
RRID:AB_10708816
IB(1:2500)
AntibodyRabbit monoclonal anti-tubulinCell Signaling TechnologyCat# 9099, RRID:AB_10695471IB(1:2000)
AntibodyRabbit polyclonal anti-LET-418Novus BiologicalsCat# 48960002, RRID:AB_10708820IB(1:1000)
AntibodyMouse monoclonal anti-SMO-1Pelisch et al., 2014Gift from Ronald T HayIB(1:500) freshly purified from hybridoma cell culture
AntibodyRabbit polyclonal anti-ubiquitinAbcamCat# ab7780; RRID:AB_306069IB(1:1000)
AntibodyRabbit polyclonal anti-HRDE-1/WAGO-9Ashe et al., 2012Gift from Eric A MiskaIB(1:500)
AntibodyGoat anti-mouse IgG (HRP-conjugated)Thermo Fisher ScientificCat# 62-6520; RRID:AB_2533947IB(1:2500)
AntibodyMouse Anti-rabbit IgG light (HRP-conjugated)AbcamCat# ab99697; RRID:AB_10673897IB(1:3000)
AntibodyMouse monoclonal anti-histone H3, di methyl K9AbcamCat# ab1220;
RRID:AB_449854
IF(1:100)
AntibodyMouse monoclonal anti-histone H3, acetyl K9AbcamCat# ab12179; RRID:AB_298910IF(1:100)
AntibodyRabbit polyclonal anti-trimethyl histone H3, K4MilliporeCat# 07-473; RRID:AB_1977252IF(1:100)
AntibodyGoat anti-mouse IgG (H+L) Alexa Fluor 488Thermo Fisher ScientificCat# A-11001; RRID:AB_2534069IF(1:1000)
AntibodyGoat anti-mouse IgG (H+L) Alexa Fluor 594Thermo Fisher ScientificCat# A-11005; RRID:AB_2534073IF(1:1000)
AntibodyGoat anti-rabbit IgG (H+L) Alexa Fluor 568Thermo Fisher ScientificCat# A-11011; RRID:AB_143157IF(1:1000)
AntibodyMouse monoclonal anti-FLAGSigma-AldrichCat# F3165; RRID:AB_259529For CHIP
AntibodyMouse monoclonal anti-dimethyl histone H3, K9MBL internationalCat# MABI0307; RRID:AB_11124951For CHIP
AntibodyRabbit polyclonal anti-histone H3, trimethyl K9MilliporeCat# 07-523; RRID:AB_310687For CHIP
AntibodyRabbit polyclonal anti-histone H3, acetyl K9AbcamCat# ab4441; RRID:AB_2118292For CHIP
Strain, strain backgroundC. elegans strainsThis studySupplementary file 3
Strain, strain backgroundE. coli: Strain OP50Caenorhabditis Genetics CenterWormBase: OP50
Strain, strain backgroundE. coli: Strain HT115Caenorhabditis Genetics CenterWormBase: HT115
Strain, strain backgroundE. coli: Ahringer collectionLaboratory of C. MelloN/A
Peptide,
recombinant protein
Ex Taq DNA polymeraseTakaraCat# RR001C
Peptide,
recombinant protein
iProof high fidelity DNA polymeraseBio-RadCat#1725302
Peptide,
recombinant protein
Alt-R S.p. Cas9 Nuclease V3Integrated DNA Technologies (IDT)Cat# 1081058CRISPR reagent
Peptide,
recombinant protein
Alt-R A.s. Cas12a (Cpf1) V3Integrated DNA Technologies (IDT)Cat# 1081068CRISPR reagent
Peptide,
recombinant protein
GFP-binding protein (GBP) beadsHomemadeN/A
Peptide, recombinant proteinHybridase Thermostable RNase HLucigen CorporationCat# H39500
Peptide, recombinant proteinTurbo DNaseThermo Fisher ScientificCat# AM2238
Peptide, recombinant proteinSuper Script III Reverse TranscriptaseThermo Fisher ScientificCat# 18080085
Peptide, recombinant proteinT4 RNA ligase 1New England BiolabsCat# M0437M
Peptide, recombinant proteinT4 RNA ligase 2New England BiolabsCat# M0242L
Peptide, recombinant proteinSuper Script III Reverse TranscriptaseThermo Fisher ScientificCat# 18080085
Chemical compound, drugEthidium bromideSigma-AldrichCat# E1510
Chemical compound, drugIsopropyl-β -D-thiogalactoside (IPTG)Sigma-AldrichCat# 11411446001
Chemical compound, drugAmpicillinSigma-AldrichCat# A9518
Chemical compound, drugTetracyclineSigma-AldrichCat# 87128
Chemical compound, drugIndole-3-acetic acid (IAA)Alfa AesarCat# A10556
Chemical compound, drugTetramisole hydrochlorideSigma-AldrichCat# L9756-5G
Chemical compound, drugParaformaldehyde 16% solutionElectron Microscopy ScienceCat# Nm15710
Chemical compound, drugFormaldehyde, 36.5–38% in H2OSigma-AldrichCat# F8775
Chemical compound, drugPBSLife TechnologiesCat# AM9615
Chemical compound, drugTween20Fisher BioReagentsCat# BP337-500
Chemical compound, drugBovine serum albumin (BSA)Life TechnologiesCat# AM2618
Chemical compound, drug1M HEPES, pH7.4TEKnovaCat# H1030
Chemical compound, drugSodium citrate dihydrateThermo Fisher ScientificCat# BP337500
Chemical compound, drugTriton X-100Sigma-AldrichCat# T8787-250ml
Chemical compound, drugcOmplete EDTA-free Protease Inhibitor CocktailRocheCat# 11836170001
Chemical compound, drugNP-40EMD MilliporeCat# 492018
Chemical compound, drugTris (Base)AvantorCat# 4099–06
Chemical compound, drugEthylenediaminetetraacetic acid disodium salt dihydrateSigma-AldrichCat# E1644
Chemical compound, drugTE buffer, pH 8.0Thermo Fisher ScientificCat# AM9858
Chemical compound, drugSodium dodecyl sulfate (SDS)Sigma-AldrichCat# L3771-100G
Chemical compound, drugSodium chloride (NaCl)Genesee ScientificCat# 18-214
Chemical compound, drugMagnesium chloride (MgCl2)Sigma-AldrichCat# M8266
Chemical compound, drugDL-Dithiothreitol (DTT)Sigma-AldrichCat# D0632-10G
Chemical compound, drugCalcium chloride (CaCl2)Sigma-AldrichCat# C5080
Chemical compound, drugPotassium chloride (KCl)Sigma-AldrichCat# P9541
Chemical compound, drugGuanidine-HClSigma-AldrichCat# G3272
Chemical compound, drugImidazoleSigma-AldrichCat# 792527
Chemical compound, drugβ-MercaptoethanolSigma-AldrichCat# M6250
Chemical compound, drugSodium phosphate, dibasicSigma-AldrichCat# S7907
Chemical compound, drugSodium phosphate, monobasicSigma-AldrichCat# S0751
Chemical compound, drugUreaThermo Fisher ScientificCat# Ac327380010
Chemical compound, drugTrichloroacetic acid (TCA)Sigma-AldrichCat# T0699
Chemical compound, drug1-Bromo-3-chloropropaneSigma-AldrichCat# B9673
Chemical compound, drugGlycineThermo Fisher ScientificCat# BP381-1
Chemical compound, drugTRI reagentSigma-AldrichCat# T9424
Chemical compound, drugTrypsinNew England BiolabsCat# P8101S
Commercial assay, kitSlowFade Diamond antifade Mountant with DAPILife TechnologiesCat# S36964
Commercial assay, kitQuick start Bradford 1xdye reagentBio-RadCat# 5000205
Commercial assay, kitNuPAGE LDS sample buffer (4x)Thermo Fisher ScientificCat# NP0008
Commercial assay, kitGlycoBlue CoprecipitantThermo Fisher ScientificCat# AM9515
Commercial assay, kitNi-NTA resinQiagenCat# 30210
Commercial assay, kitpCR-Blunt II Topo cloning kitThermo Fisher ScientificCat# K280020
Commercial assay, kitMinElute PCR purification KitQiagenCat# 28006
Commercial assay, kitChIP DNA clean and concentrator KitZymo ResearchCat# 5205
Commercial assay, kitProteoSilver Plus Silver Stain KitSigma-AldrichCat# PROT-SIL2
Commercial assay, kitTrans-blot Turbo Mini NC Transfer PacksBio-RadCat# 1704158
Commercial assay, kitLumi-Light Plus western blotting substrateSigma-AldrichCat# 12015196001
Commercial assay, kitHyperfilm ECLThermo Fisher ScientificCat# 45001507
Commercial assay, kitmirVana miRNA Isolation KitThermo Fisher ScientificCat# AM1561
Commercial assay, kitKAPA RNA HyperPrep with RiboErase (KK8560)RocheCat# 08098131702
Commercial assay, kitKAPA single-indexed adapter kit (KK8700)RocheCat# 08005699001
Commercial assay, kitChIP-Grade Protein A/G Magnetic BeadsThermo Fisher ScientificCat# 26162
Commercial assay, kitIllumina NextSeq 500/550 v2.5 kit (75 cycles)Illumina, Inc.Cat# 20024906
Commercial assay, kitIllumina NextSeq 500/550 v2.5 kit (150 cycles)Illumina, Inc.Cat# 20024907
Recombinant DNA reagentPeft3::cas9 vector (backbone: blunt II topo vector)Friedland et al., 2013N/ABackbone: blunt II topo vector (Kim et al., 2021)
Recombinant DNA reagentpRF4: injection marker, rol-6(su1006)Mello et al., 1991N/ABackbone: blunt II topo vector (Kim et al., 2021)
Recombinant DNA reagentsmo-1 sgRNA plasmidThis studySupplementary file 4
Recombinant DNA reagentmep-1 sgRNA plasmidKim et al., 2021Supplementary file 4
Sequence-based reagentList of gRNA sequencesThis studySupplementary file 4
Sequence-based reagentAlt-R CRISPR-Cas9 tracrRNAIntegrated DNA Technologies (IDT)Cat# 1072534CRISPR reagent
Sequence-based reagentAnti-rRNA Oligos for C. elegansThis study (homemade)N/A
Software, algorithmGraphPad Prism version 8.2.1GraphPad Softwarehttp://www.graphpad.com
Software, algorithmSalmonPatro et al., 2017v1.1.0
Software, algorithmDEseq2Love et al., 2014v1.26.0
Software, algorithmBowtie2Langmead and Salzberg, 2012v2.2.3
Software, algorithmPicard-toolsBroad Institute, 2019v1.131
Software, algorithmMACS2Feng et al., 2012v2.1.1.20160309
Software, algorithmBedToolsQuinlan and Hall, 2010v2.25.0
Software, algorithmIGVRobinson et al., 2017v2.7.2

Data availability

RNA Sequencing Data have been deposited in SRA under accession codes Bioproject: PRJNA657279. ChIP Sequencing Data have been deposited in SRA under accession codes Bioproject: PRJNA657194. All data generated or analyzed during this study are included in the manuscript and supplementary files.

The following data sets were generated
    1. Mello CC
    (2021) NCBI BioProject
    ID PRJNA657279. RNA seq of NuRD complex mutants and piRNA pathway mutants.
    1. Mello CC
    (2021) NCBI BioProject
    ID PRJNA657194. ChIP seq of NuRD complex components and histone modifications.

References

  1. Book
    1. Askjaer P
    2. Ercan S
    3. Meister P
    (2014)
    Modern Techniques for the Analysis of Chromatin and Nuclear Organization in C. Elegans
    WormBook.
    1. Brenner S
    (1974)
    The genetics of Caenorhabditis elegans
    Genetics 77:71–94.
    1. Chen Z
    2. Han M
    (2001)
    Role of C. elegans lin-40 MTA in vulval fate specification and morphogenesis
    Development 128:4911–4921.
    1. Partridge JF
    2. Borgstrøm B
    3. Allshire RC
    (2000)
    Distinct protein interaction domains and protein spreading in a complex centromere
    Genes & Development 14:783–791.

Decision letter

  1. Tony Hunter
    Reviewing Editor; Salk Institute for Biological Studies, United States
  2. Kevin Struhl
    Senior Editor; Harvard Medical School, United States
  3. Tony Hunter
    Reviewer; Salk Institute for Biological Studies, United States
  4. Federico Pelisch
    Reviewer; University of Dundee, United Kingdom

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

Acceptance summary:

These studies reveal that sumoylation of HDA-1, a type 1 histone deacetylase (HDAC), plays a key role in establishing transcriptional silencing of piRNA-regulated genes in C. elegans. Genetic analysis provides strong evidence that the SUMO pathway is involved in piRNA silencing, and further mechanistic analysis demonstrates this involves sumoylation of two lysines in the tail of HDA-1. HDA-1 sumoylation promotes its association with the NuRD chromatin modifier complex, which enhances local H3K9ac deacetylation, resulting in negative regulation of hundreds of target genes that is instrumental in the inherited RNAi pathway.

Decision letter after peer review:

Thank you for submitting your article "HDAC1 SUMOylation promotes Argonaute directed transcriptional silencing in C. elegans" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, including Tony Hunter as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Kevin Struhl as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Federico Pelisch (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

As the editors have judged that your manuscript is of interest, but as described below that additional experiments are required before it is published, we would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Summary:

In this paper, your studies showed that sumoylation of HDA-1, a type 1 HDAC, at two C-terminal Lys residues plays a role in establishing transcriptional silencing of piRNA-regulated genes in C. elegans through enhanced NuRD complex interaction and histone H3 deacetylation. The reviewers all found the link between HDA-1 sumoylation and silencing to be interesting, but raised a number of issues that need to be addressed. In particular, more mechanistic insights into

Essential revisions:

1. The evidence that sumoylation of HDA-1 increases its silencing function is reasonably strong, but no direct evidence is provided that HDA-1 sumoylation at K444 and K459 increases its deacetylase activity, as is proposed. For this purpose, biochemical assays comparing unmodified HDA-1 (desumoylated WT or KKRR mutant HDA-1) with sumoylated HDA-1 (or SUMO-fused HDA-1) to measure changes in deacetylatase activity are required. In this connection, there are also no mechanistic insights into how the presence of a SUMO moiety at either K444 or K459 site in the C-terminal tail would increase HDA-1 catalytic activity. Finally, your evidence for sumoylation is based on K to R mutations of these two Lys, and direct mass spec evidence that the K444 and K459 sites are in fact sumoylated in vivo would be desirable, although not essential.

2. Although the evidence that sumoylated HDA-1 interacts with the NuRD complex is reasonable, it remains unclear how sumoylation of HDA-1 increases its recognition by the NuRD complex. To determine whether the HDA-1 SUMO moieties are directly involved in the interaction, testing for SIMs (and/or UIMs) in MEP-1, or another subunit of the NuRD complex subunit is needed, followed by verification of the importance of these residues in vivo or through in vitro interaction assays.

3. Evidence is provided that HDA-1 sumoylation decreases total H3K9ac histone acetylation in vivo, but the use of ChIP-seq to test if HDA-1 and NuRD complex recruitment, and local histone H3 acetylation are affected at known target genes in the KKRR mutant and HDA-1 degron strains versus the wild type strain, with correlated RNA-seq data in the same lines, would significantly strengthen the conclusion that HDA-1 sumoylation is required for piRNA regulation. The addition of ChIP-seq data for HDA-1 and HDA-1-KKRR genome-wide distribution profiles would further strengthen the authors' claims, but is not essential.

4. A possible role of the GEI-17 E3 SUMO ligase in the regulation of HDA-1 and piRNA-regulated gene silencing was excluded through the use of a gei-17 mutant. However, while neither gei-17 nor pie-1 Individually affected piRNA reporter expression in P0, the authors need to test a double mutant to confirm a lack of synergy.

5. Based on the data presented only a small fraction of the HDA-1 population was sumoylated, and the authors need discuss how sumoylation of such a small fraction of the HDA-1 protein population would be able to exert the functional effects on HDA-1 activity they observed.

6. The authors could do a more comprehensive job of reviewing the relevant literature on the role of sumoylation in Piwi-mediated silencing in other organisms, e.g. Ninova et al. Mol Cell 77:556, 2020.

Additional points are raised in the individual reviews below and could be considered, if the authors believe that addressing them would strengthen the manuscript.

Reviewer #1:

The evidence that sumoylation of HDA-1, a type 1 HDAC, plays a key role in establishing transcriptional silencing of piRNA-regulated genes in C. elegans is quite convincing. The genetic analysis demonstrating that the SUMO pathway is involved in piRNA silencing is strong, and the mutational evidence that this involves sumoylation of two Lys in the tail of HDA-1 is reasonable. Likewise, the finding that HDA-1 sumoylation promotes association with NuRD complex components and association of MEP-1, an HDA-1 interactor, with chromatin regulators is convincing. In addition, the evidence that HDA-1 sumoylation increases H3K9ac deacetylation in vivo, leading to negative regulation of hundreds of target genes, and plays a role in the inherited RNAi pathway is solid.

While the overall conclusion provides an interesting advance in understanding mechanisms of piRNA-mediated gene silencing in C. elegans, the paper is lacking any biochemical analysis of the effects of sumoylation on HDA-1 activity and its association with other transcriptional regulators.

1. The authors mapped two sumoylation sites close to the C terminus of HDA-1, K444 and K459, based on extremely weak homology with two established sumoylation sites in human HDAC1 that are reported to be important for transcriptional repression (N.B. the authors should indicate here that David et al. reported that K444/476R HDAC1 had reduced transcriptional repression activity in reporter assays.). While the two human sites conform to the sumoylation site consensus, ψKXE, neither K444 nor K459 in HDA-1 fits this consensus (possibly one could argue that K444 is in an inverted motif). The fact that the KKRR mutant HDA-1 is no longer sumoylated is consistent with these two Lys being sumoylated, but it would be reassuring to have direct MS evidence that K444 and K459 are indeed sumoylated, which could be achieved using a SUMO Thr91Arg mutant that generates a GlyGly stub upon trypsin digestion, among other methods.

2. It remains unclear how sumoylated HDA-1 is recognized by MEP-1 for assembly into the NuRD complex. Does MEP-1, or another NuRD subunit, have a SIM that could facilitate direct interaction of MEP-1 and sumoylated HDA-1?

3. As the authors discuss, it is surprising that the HDA-1(KKRR)::SUMO protein, which in effect is a constitutively sumoylated form of HDA-1 that will interact constitutively with MEP-1/NuRD, does not have more deleterious effects on the organism, since according to the data in Figure 2B, the stoichiometry of endogenous HDA-1 sumoylation was extremely low. Of course, low sumoylation stoichiometry, which is a general issue with sumoylation studies, means that only a very small fraction of the HDA-1 endogenous population will be able to engage with the silencing complexes at any one time. This point is also worth discussion.

4. Page 5: Here, and elsewhere, the authors claim that sumoylation of the two C-terminal Lys activates HDA-1 histone deacetylase activity, but provide no direct evidence for this statement. There are no HDAC assays, and it is unclear how C-terminal SUMO residues distant from the catalytic domain would alter its enzymatic activity, unless there is a SIM motif in HDA-1 that might allow for intramolecular interaction with SUMO residues at the tail leading to a conformation change. Did the authors check for a SIM motif in HDA-1? The fact that adding SUMO to the C-terminus rather than one or both of the two Lys would also have to be taken into account in determining bow sumoylation might "activate" HDA-1. To demonstrate that sumoylation activated HAD-1 in vitro deacetylation assays would need to be carried out comparing the activities of unmodified and sumoylated HDA-1. Instead of enzymatic activation, it is possible that the PIE-1 interaction and HDA-1 sumoylation results in relocalization of HDA-1 within the nucleus to facilitate more efficient H3K9ac deacetylation.

Reviewer #2:

In their manuscript, Kim et al. describe a role for HDAC1 (HDA-1) sumoylation in Argonaute-directed transcriptional silencing. The authors suggest that sumoylation of HDA-1 is important for proper assembly of the NuRD deacetylase complex. The role of SUMO modification in heterochromatin has been extensively documented and it is a very interesting topic. The current manuscript provides a very interesting set of results on this topic. I have list of comments, suggestions, questions, and concerns, which are listed below, especially related to the first half of the results:

1. A general question would be how can HDA-1 sumoylation, which is barely detectable, account for such a big 'positive' effect on complex assembly? HDA-1 SUMO modification seems around 10% after enriching for SUMO-modified proteins, which means that stoichiometry will be way lower than this. While this is common for SUMO-modified proteins, it does make it difficult to associate with a 'simple' model.

2. In Figure 1, a schematic of the sensor used throughout the study would benefit the reader.

3. In Figure 1, have the authors checked if the 10xHis::tagged smo-1 has the same effect as the 3xflag::smo-1 (i.e. is it also a partial loss of function allele)?

4. In Figure 1 it would be nice to see the global SUMO conjugation levels in the different conditions, particularly in the smo-1(RNAi), 3xflag::smo-1, and ubc-9(G56R).

5. Also Figure 1, was gei-17 depletion/deletion checked in any way (i.e. WB)? Did the authors consider other SUMO E3 ligase, such as the mms-21 orthologue?

6. While I am not a big fan of fusing SUMO to proteins, in this case it seems like a very reasonable thing to do, considering the modification sites are located very close to the C-terminal end of the protein. Did the authors check an N-terminal fusion?

7. In Figure 2B, it becomes very clear that the level of SUMO modification of HDA-1 is extremely small, barely detectable after an enrichment method. I also wonder why the gels were cropped so tightly, especially considering that in Figure 3 there is an additional band corresponding to ubiquitylated, sumoylated HDA-1. in vitro modification assays would be helpful. HDA-1 alongside a known and characterised SUMO substrate would indicate how good a substrate HDA-1 is.

8. In Figure 2D, is the difference between HDA-1(KKRR)::SUMO and HDA-1::SUMO significant?

9. In Figure 3A-C, it would be useful to control whether the GFP::HDA-1 fusion behaves as the untagged one in the sensor assay (wt vs. KKRR).

10. I have a few questions regarding Figure 3D:

i. Considering the extremely low level of HDA-1 sumoylation, did the authors detect SUMO and ub conjugated HAD-1 (not the SUMO usion)?

ii. Is ub conjugated to SUMO or to HDA-1?

iii. Does MEP-1 contain any obvious SIMs and or UIMs?

iv. To make a stronger case for the SUMO-dependent interaction model, in vitro interaction assays with recombinant proteins would be extremely useful.

11. In the discussion, the authors compare the lack of requirement for GEI-17 in their manuscript with the requirement for Su(var)2-10 in flies. It is very important to back this claim that the authors control GEI-17 depletion (as pointed out in 5).

Overall, I think this manuscript provides a very interesting set of results and I believe that, with the addition of some simple biochemical experiments, the quality and impact of the overall work would be much greater.

Reviewer #3:

This manuscript by Kim et al. describes a role of SUMOylation in Argonaute-directed transcriptional silencing in C. elegans. The Authors found that SUMOylation of the histone deacetylase HDA-1 promotes its interaction with both the Argonatue target recognition complex as well as the chromatin remodeling NuRD complex. This enables initiation of target silencing. Impaired SUMOylation of HDA-1 leads to loss of interactions with several protein complexes, reduced silencing of piRNA targets, and reduced brood size. While the findings and claims are interesting, some of the novelty is overemphasized and some of the claims are not fully supported by the data.

1. The importance of HDA-1 SUMOylation for transcriptional repression. The title "HDAC1 SUMOylation promotes Argonaute directed transcriptional silencing in C. elegans" implies a central role of SUMOylation in piRNA-mediated transcriptional silencing. The Argonaute HRDE-1/WAGO-9 targets countless transposons as shown previously and also in this manuscript (Figure S3), and so do the HDA-1 degron and Ubc9 mutant, indicating that histone deacetylation and protein SUMOylation are essential processes in TE silencing. However, the HDA-1 SUMOylation mutant (KKRR) only slightly affects 6 TE families (Figure S3), indicating that SUMOylation of HDA-1 might not be a key mediator of this process. Furthermore, the authors write that "Our findings suggest how SUMOylation of HDAC1 promotes the recruitment and assembly of an Argonaute-guided chromatin remodeling complex to orchestrate de novo gene silencing in the C. elegans germline.", but then they also state that "Comparison with mRNA sequencing data from auxin-treated degron::hda-1 animals revealed an even more extensive overlap with Piwi pathway mutants (Figure S2B), indicating that HDA-1 also promotes target silencing independently of HDA-1 SUMOylation." Based on their results and their own interpretations, I find that the importance of HDA-1 SUMOylation in piRNA-dependent transcriptional silencing is overemphasized.

Additionally, the model (Figure 7) implies that for initiation of silencing WAGO recruits HDA-1 to targets. This should be tested by analyzing HDA-1 distribution over WAGO targets in WT and upon loss of WAGO.

2. The mechanistic role of HDA-1 SUMOylation.

On page 17 (amongst other places) the authors claim that "The SUMOylation of HDA-1 promotes its activity, while also promoting physical interactions with other components of a germline nucleosome-remodeling histone deacetylase (NuRD) complex, as well as the nuclear Argonaute HRDE-1/WAGO-9 and the heterochromatin protein HPL-2 (HP1)".

• Regarding activity: Loss of deacetylation/silencing in the SUMO mutant might be due to loss of enzymatic activity, but it might also be due to defects in recruitment/complex formation. There is no data that proves altered enzymatic activity. In fact, Figure 6 indicates SUMO-dependent interaction of WAGO-9 with HDA-1, implying that recruitment is affected. To distinguish between activity and recruitment, at the very least, the authors would need to show that HDA-1 localization to its genomic targets is unaltered upon mutating its SUMOylation site (ChIP-seq of wt and KKRR mutant), while H3K9ac is increased (K9ac ChIP-seq in wt and KKRR mutant) in the mutant. This, in combination with HDA-1 localization in wt and WAGO-9 loss would imply whether complex formation to recruit HDA-1 or HDA-1 enzymatic activity is mostly affected by SUMOylation.

• Regarding physical interactions: Figure 3D shows that if we fuse a SUMO residue to HDA-1, it will interact with MEP-1, while SUMOylation deficient HDA-1 mutant doesn't interact. However, for the WT HDA-1 control, we only see unSUMOylated protein interacting with MEP-1. Furthermore, in the MEP-1 IPs of samples that should contain SUMO-fused HDA-1, the authors detect a lot of "cleaved", unSUMOylated HDA-1. Unless cleavage happened after IP, during elution (unlikely, and there is "cleaved" HDA-1 in the inputs), these findings argue that the interaction with MEP-1 is not mediated by HDA-1 SUMOylation. An interaction between MEP-1 and unmodified HDA-1 is also shown in the accompanying manuscript, which appears to be dependent on Pie-1 SUMOylation. Thus, SUMOylation of HDA-1 alone seems unlikely to be the major factor necessary for silencing complex assembly. (as a side question: Does the protease inhibitor cocktail used inhibit de-SUMOylation enzymes? I am concerned that deSUMOylating enzymes might compromise some result interpretations.)

• Regarding functional relevance of HDA-1 acetylation: On pages 12/13 authors claim that because "HDA-1(KKRR) animals and mep-1-depleted worms revealed dramatically higher levels of H3K9Ac compared to wild-type" and "HDA-1, LET-418/Mi-2, and MEP-1 bind heterochromatic", "SUMOylation of HDA-1 appears to drive formation or maintenance of germline heterochromatin regions of the genome." These correlations do not prove function. The authors have performed H3K9me2 (although not H3K9-ac) ChIP-seq in WT, KKRR mutant and HDA-1 degron worms, yet do not analyze globally whether acetylation is lost on genes that are affected (change in RNA-seq vs. change in K9me2 or acetyl). To support the claim that SUMOylation of HDA-1 drives deacetylation and heterochromatin formation, it would be important to show changes in H3K9Ac levels (or other acetyl marks) and potentially NuRD component occupancy between control and HDA-1 SUMOylation-deficient animals at specific targets (i.e. genes derepressed upon loss of SUMOylation identified in RNA-seq, and the reporter locus).

3. The authors claim (p17) that "initiation of transcriptional silencing requires SUMOylation of conserved C-terminal lysine residues in the type-1 histone deacetylase HDA-1". I do not see any supporting data that has separately looked at formation/initiation and maintenance of silencing (a technically challenging experiment).

4. The authors repeatedly claim that gei-17 does not play a role in piRNA target silencing, based on loss of gei-17 not affecting the piRNA reporter (Figure 1B). At the same time, they claim that pie-1 plays a role, even though it likewise does not affect the piRNA reporter (it affects the reporter only in F3; data on gei-17 effect in F3 is not present). In the accompanying paper, the authors show that while gei-17 loss by itself causes only moderate effect on extra intestine cells, combined with Pie-1 loss the effect is more severe than when Pie-1 loss is combined with Ubc9 or smo loss. This to me indicates an important role of gei-17 in inhibiting differentiation of germline stem cells to somatic tissues, but these effects are likely synergistic and thus masked by Pie-1. Individually neither Gei-17 nor Pie-1 show an effect on piRNA reporter in P0, but to confirm lack of synergy, their effects of should be tested together. Although possible, the present data is insufficient to rule of gei-17 involvement.

https://doi.org/10.7554/eLife.63299.sa1

Author response

Essential revisions:

1. The evidence that sumoylation of HDA-1 increases its silencing function is reasonably strong, but no direct evidence is provided that HDA-1 sumoylation at K444 and K459 increases its deacetylase activity, as is proposed. For this purpose, biochemical assays comparing unmodified HDA-1 (desumoylated WT or KKRR mutant HDA-1) with sumoylated HDA-1 (or SUMO-fused HDA-1) to measure changes in deacetylatase activity are required. In this connection, there are also no mechanistic insights into how the presence of a SUMO moiety at either K444 or K459 site in the C-terminal tail would increase HDA-1 catalytic activity.

As noted in our reply to the PIE-1 paper, we did not mean to imply that SUMO modification of HDAC increases its enzymatic activity. This possibility had not even occurred to us and seems very unlikely. We now emphasize the genetic and molecular genetic studies as support for the importance of direct HDAC SUMOylation in Argonaute-mediated germline surveillance. We have opted not to attempt in vitro HDAC assays, as it is not clear to us what additional insights any unlikely changes in activity might have. Moreover, such a study would very likely require structural investigations of any changes to the protein active site. We have now made it clear throughout the paper that the SUMO modifications are thought to alter HDAC activity indirectly by regulating its association with co-factors.

Finally, your evidence for sumoylation is based on K to R mutations of these two Lys, and direct mass spec evidence that the K444 and K459 sites are in fact sumoylated in vivo would be desirable, although not essential.

Unfortunately, MS detection of SUMOylated sites requires a mutation in smo-1 that is lethal in worms. We mention this negative result in the PIE-1 paper.

2. Although the evidence that sumoylated HDA-1 interacts with the NuRD complex is reasonable, it remains unclear how sumoylation of HDA-1 increases its recognition by the NuRD complex. To determine whether the HDA-1 SUMO moieties are directly involved in the interaction, testing for SIMs (and/or UIMs) in MEP-1, or another subunit of the NuRD complex subunit is needed, followed by verification of the importance of these residues in vivo or through in vitro interaction assays.

Thank you for raising this interesting point regarding SIMs in MEP-1. Our results clearly show that HDA-1-SUMO promotes its interactions with the NuRD complex. Whether or not it does this through the SIMs in MEP-1 or another NuRD complex factor is not relevant to our model or story. We have generated and characterized double-SIM mutants of MEP-1. Interestingly, the MEP-1 double-SIM mutant shows a strong desilencing of the piRNA sensor, but the worms are sterile due to an oogenesis defect—like the mep-1 null mutant. Interpreting this phenotype is complicated by the likelihood that the SIMs on MEP-1 interact with not only HDA-1 but also other SUMOylated proteins. Future studies will have to address this important detail.

The issue of how HDA-1 SUMoylation promotes its interactions in the adult germline are now directly discussed as follows:

“Paradoxically in the embryo HDA-1 is not SUMOylated and yet interacts robustly with MEP-1.[…] Thus although the SIM motifs in MEP-1 may be important for its interaction with HDA-1-SUMO they may also be required for other functions or for interactions with other essential co-factors.”

3. Evidence is provided that HDA-1 sumoylation decreases total H3K9ac histone acetylation in vivo, but the use of ChIP-seq to test if HDA-1 and NuRD complex recruitment, and local histone H3 acetylation are affected at known target genes in the KKRR mutant and HDA-1 degron strains versus the wild type strain, with correlated RNA-seq data in the same lines, would significantly strengthen the conclusion that HDA-1 sumoylation is required for piRNA regulation. The addition of ChIP-seq data for HDA-1 and HDA-1-KKRR genome-wide distribution profiles would further strengthen the authors' claims, but is not essential.

The genetic evidence that HDA-1 SUMOylation promotes Argonaute-mediated silencing is quite strong, and is supported by our RNA-seq data on dissected gonads and by the heritable silencing defect of HDA-1(KKRR) worms in response to a dsRNA trigger. ChIP-seq data would be nice to have, but would require purifying germline nuclei away from the somatic tissues of adults—a technical challenge. A few dozen dissected gonads would yield insufficient material for ChIP-seq experiments.

4. A possible role of the GEI-17 E3 SUMO ligase in the regulation of HDA-1 and piRNA-regulated gene silencing was excluded through the use of a gei-17 mutant. However, while neither gei-17 nor pie-1 Individually affected piRNA reporter expression in P0, the authors need to test a double mutant to confirm a lack of synergy.

Thank you for this excellent suggestion. Indeed, the double shows a very strong de-silencing phenotype as indicated in Figure 6 of the PIE-1 paper. We had trouble making this double due to a haploinsufficiency, but finally succeeded using CRISPR.

5. Based on the data presented only a small fraction of the HDA-1 population was sumoylated, and the authors need discuss how sumoylation of such a small fraction of the HDA-1 protein population would be able to exert the functional effects on HDA-1 activity they observed.

We apologize for neglecting to discuss this important point. The second section of the discussion now addresses this issue at some length:

SUMO, a potent genetic modifier with an elusive biochemical signature

Given the strong genetic evidence that the modification of HDA-1 by SUMO promotes piRNA silencing in the adult germline, we were surprised that the conjugated isoform of HDA-1 was undetectable in our IP assays from adult animals. […] Consistent with this idea, an HDA-1::SMO-1 translational fusion protein with an abnormal linkage via the N-terminus of SUMO was stable in protein lysates, strongly rescued the silencing defects of the presumptive SUMO-acceptor mutant protein HDA-1(KKRR), and dramatically enhanced the detection of protein-protein interactions between HDA-1 and components of an adult-stage NuRD complex.”

6. The authors could do a more comprehensive job of reviewing the relevant literature on the role of sumoylation in Piwi-mediated silencing in other organisms, e.g. Ninova et al. Mol Cell 77:556, 2020.

We agree. We now review the literature more comprehensively in the final section of the discussion as follows:

Parallels in the role of SUMOylation in Piwi silencing in insects, mice and worms

Histone deacetylation is a necessary step in de novo transcriptional silencing. Yet, precisely how nuclear Argonautes orchestrate both deacetylation and the subsequent installation of silencing marks on target chromatin is not known. […]It will be interesting in the future to learn if mammalian and insect Piwi Argonautes target HDAC1 SUMOylation to promote de novo Piwi silencing”.

Additional points are raised in the individual reviews below and could be considered, if the authors believe that addressing them would strengthen the manuscript.

Reviewer #1:

The evidence that sumoylation of HDA-1, a type 1 HDAC, plays a key role in establishing transcriptional silencing of piRNA-regulated genes in C. elegans is quite convincing. The genetic analysis demonstrating that the SUMO pathway is involved in piRNA silencing is strong, and the mutational evidence that this involves sumoylation of two Lys in the tail of HDA-1 is reasonable. Likewise, the finding that HDA-1 sumoylation promotes association with NuRD complex components and association of MEP-1, an HDA-1 interactor, with chromatin regulators is convincing. In addition, the evidence that HDA-1 sumoylation increases H3K9ac deacetylation in vivo, leading to negative regulation of hundreds of target genes, and plays a role in the inherited RNAi pathway is solid.

While the overall conclusion provides an interesting advance in understanding mechanisms of piRNA-mediated gene silencing in C. elegans, the paper is lacking any biochemical analysis of the effects of sumoylation on HDA-1 activity and its association with other transcriptional regulators.

1. The authors mapped two sumoylation sites close to the C terminus of HDA-1, K444 and K459, based on extremely weak homology with two established sumoylation sites in human HDAC1 that are reported to be important for transcriptional repression (N.B. the authors should indicate here that David et al. reported that K444/476R HDAC1 had reduced transcriptional repression activity in reporter assays.).

We thank the reviewer for raising this point. We have added to the discussion on this interesting vertebrate work.

While the two human sites conform to the sumoylation site consensus, ψKXE, neither K444 nor K459 in HDA-1 fits this consensus (possibly one could argue that K444 is in an inverted motif). The fact that the KKRR mutant HDA-1 is no longer sumoylated is consistent with these two Lys being sumoylated, but it would be reassuring to have direct MS evidence that K444 and K459 are indeed sumoylated, which could be achieved using a SUMO Thr91Arg mutant that generates a GlyGly stub upon trypsin digestion, among other methods.

As noted above and in the PIE-1 paper, we attempted these experiments. However, mass spec detection of SUMO-modified peptides requires a smo-1 L88K mutant, which is inviable.

2. It remains unclear how sumoylated HDA-1 is recognized by MEP-1 for assembly into the NuRD complex. Does MEP-1, or another NuRD subunit, have a SIM that could facilitate direct interaction of MEP-1 and sumoylated HDA-1?

This point is now discussed and was addressed in the earlier comments above.

3. As the authors discuss, it is surprising that the HDA-1(KKRR)::SUMO protein, which in effect is a constitutively sumoylated form of HDA-1 that will interact constitutively with MEP-1/NuRD, does not have more deleterious effects on the organism, since according to the data in Figure 2B, the stoichiometry of endogenous HDA-1 sumoylation was extremely low. Of course, low sumoylation stoichiometry, which is a general issue with sumoylation studies, means that only a very small fraction of the HDA-1 endogenous population will be able to engage with the silencing complexes at any one time. This point is also worth discussion.

This too was addressed above.

4. Page 5: Here, and elsewhere, the authors claim that sumoylation of the two C-terminal Lys activates HDA-1 histone deacetylase activity, but provide no direct evidence for this statement. There are no HDAC assays, and it is unclear how C-terminal SUMO residues distant from the catalytic domain would alter its enzymatic activity, unless there is a SIM motif in HDA-1 that might allow for intramolecular interaction with SUMO residues at the tail leading to a conformation change. Did the authors check for a SIM motif in HDA-1? The fact that adding SUMO to the C-terminus rather than one or both of the two Lys would also have to be taken into account in determining bow sumoylation might "activate" HDA-1. To demonstrate that sumoylation activated HAD-1 in vitro deacetylation assays would need to be carried out comparing the activities of unmodified and sumoylated HDA-1. Instead of enzymatic activation, it is possible that the PIE-1 interaction and HDA-1 sumoylation results in relocalization of HDA-1 within the nucleus to facilitate more efficient H3K9ac deacetylation.

We apologize again for our careless use of the word “activates.” As noted above, we have changed the manuscript to correct this. It is very unlikely that the enzyme itself is more active but rather that its access to substrate is increased due to interactions with the NuRD components and the Argonaute protein WAGO-9.

Reviewer #2:

In their manuscript, Kim et al. describe a role for HDAC1 (HDA-1) sumoylation in Argonaute-directed transcriptional silencing. The authors suggest that sumoylation of HDA-1 is important for proper assembly of the NuRD deacetylase complex. The role of SUMO modification in heterochromatin has been extensively documented and it is a very interesting topic. The current manuscript provides a very interesting set of results on this topic. I have list of comments, suggestions, questions, and concerns, which are listed below, especially related to the first half of the results:

1. A general question would be how can HDA-1 sumoylation, which is barely detectable, account for such a big 'positive' effect on complex assembly? HDA-1 SUMO modification seems around 10% after enriching for SUMO-modified proteins, which means that stoichiometry will be way lower than this. While this is common for SUMO-modified proteins, it does make it difficult to associate with a 'simple' model.

This issue is now a major discussion point. Thank you! We were so thrilled to even detect the modification in vivo – which is a challenge, as you know – that we neglected to comment on its elusive nature in biochemical studies. We hope the new discussion sufficiently emphasizes this point.

2. In Figure 1, a schematic of the sensor used throughout the study would benefit the reader.

We now include a schematic in Figure 1A.

3. In Figure 1, have the authors checked if the 10xHis::tagged smo-1 has the same effect as the 3xflag::smo-1 (i.e. is it also a partial loss of function allele)?

Thank you for raising this point. This strain is fully wild type. We now make this point in the PIE-1 paper.

4. In Figure 1 it would be nice to see the global SUMO conjugation levels in the different conditions, particularly in the smo-1(RNAi), 3xflag::smo-1, and ubc-9(G56R).

In Figure 3A of the PIE-1 paper, we now show that smo-1(RNAi) broadly reduces global SUMO levels.

5. Also Figure 1, was gei-17 depletion/deletion checked in any way (i.e. WB)? Did the authors consider other SUMO E3 ligase, such as the mms-21 orthologue?

Thank you for raising this point. The gei-17 genetics are addressed in the PIE-1 paper. The degron allele does not behave like a null, perhaps because TIR1 is only expressed in the germline (and so GEI-17 still functions in the soma), or because GEI-17 is not completely depleted in the germline by the auxin-inducible degron system. We could not directly measure the level of depletion of GEI-17 in the germline due to the remaining somatic GEI-17. So we removed it from the paper.

A null allele of gei-17 failed to desilence the piRNA sensor. By contrast, the double between the gei-17 null and pie-1(K68R) desilenced the sensor in 100% of isolates.

We have not explored other E3s. Given the very strong synthetic phenotype between K68R and gei-17, however, we believe that GEI-17 is the most relevant E3. GEI-17 was also identified as a PIE-1 interactor in our two-hybrid screen.

6. While I am not a big fan of fusing SUMO to proteins, in this case it seems like a very reasonable thing to do, considering the modification sites are located very close to the C-terminal end of the protein. Did the authors check an N-terminal fusion?

We have not checked the N-terminal fusion. We were surprised this worked too, and only later after doing this did we notice the paper showing that expressing HDA-1::SUMO in the mouse model for Alzheimers improved cognitive function. It seems like HDA-1 may be ideal for this approach since the lysines are so near the C-terminus.

7. In Figure 2B, it becomes very clear that the level of SUMO modification of HDA-1 is extremely small, barely detectable after an enrichment method. I also wonder why the gels were cropped so tightly, especially considering that in Figure 3 there is an additional band corresponding to ubiquitylated, sumoylated HDA-1. in vitro modification assays would be helpful. HDA-1 alongside a known and characterised SUMO substrate would indicate how good a substrate HDA-1 is.

We were only trying to conserve space. We have expanded the cropped region of the blot to show the higher molecular weight region. Ubiquitin-SUMO-HDA-1 is not detected. We worry that in vitro approaches would not be enlightening. There are so many other factors that would likely be absent from our in vitro assays. We feel that our genetic and molecular studies provide clear evidence for HDA-1 SUMOylation and its importance in vivo. Indeed, we think that the low level of modification detected make our story more interesting, not less. We suspect that the labile nature of the SUMO conjugation creates a “detection” issue that could explain the apparent absence of HDA-1 from a MEP1-Mi2 complex (called MEC) in a fly ovarian cell line. We now discuss this issue in both papers.

8. In Figure 2D, is the difference between HDA-1(KKRR)::SUMO and HDA-1::SUMO significant?

Thank you for raising this point. No, there is no significant difference between them.

WE now show this in Figure 2D.

9. In Figure 3A-C, it would be useful to control whether the GFP::HDA-1 fusion behaves as the untagged one in the sensor assay (wt vs. KKRR).

Unfortunately, our sensor is a GFP reporter, so we would need to rebuild and validate a different sensor to do this. However, the GFP tag is inserted into the endogenous hda-1 gene, which is essential. The tagged worms are viable and healthy, suggesting it is a fully functional fusion protein.

10. I have a few questions regarding Figure 3D:

i. Considering the extremely low level of HDA-1 sumoylation, did the authors detect SUMO and ub conjugated HAD-1 (not the SUMO usion)?

We detected SUMOylated HDA-1 under denaturing conditions. In IP buffer (non-denaturing), SUMO was quickly removed from proteins. We now show this in the PIE-1 paper. We did not detect Ub-modified HDA-1 in HDA-1 IPs. Like SUMO, Ubiquitin conjugation might be labile in IP buffer or the addition of Ub to the SUMO fusion could be an artifact.

ii. Is ub conjugated to SUMO or to HDA-1?

We don’t know. We discuss the possibility that SUMO is ubiquitinated because the SUMO antibody does not detect the ubiquitinated HDA-1::SMO-1, suggesting that the SUMO epitope is masked by Ub.

iii. Does MEP-1 contain any obvious SIMs and or UIMs?

Yes, MEP-1 has two consensus SIM motifs, as discussed above. We now discuss this in both papers.

iv. To make a stronger case for the SUMO-dependent interaction model, in vitro interaction assays with recombinant proteins would be extremely useful.

We are indeed interested in understanding the nature of the physical interactions—especially why SUMO is needed only in the adult. However, such a discovery would merit its own paper and would likely require months if not years of additional work. In short, we do not think SUMO alone is responsible! Some unknown factor in the adult germline may be interfering with MEP-1:HDA-1 binding. Until this factor is identified, in vitro work is unlikely to be informative. In fact, in embryos, HDA-1 is not SUMOylated AND it interacts robustly with MEP-1. So, we expect HDA-1 to interact with MEP-1 in vitro, with or without SUMO.

11. In the discussion, the authors compare the lack of requirement for GEI-17 in their manuscript with the requirement for Su(var)2-10 in flies. It is very important to back this claim that the authors control GEI-17 depletion (as pointed out in 5).

Thanks. We addressed this point above.

Reviewer #3:

This manuscript by Kim et al. describes a role of SUMOylation in Argonaute-directed transcriptional silencing in C. elegans. The Authors found that SUMOylation of the histone deacetylase HDA-1 promotes its interaction with both the Argonatue target recognition complex as well as the chromatin remodeling NuRD complex. This enables initiation of target silencing. Impaired SUMOylation of HDA-1 leads to loss of interactions with several protein complexes, reduced silencing of piRNA targets, and reduced brood size. While the findings and claims are interesting, some of the novelty is overemphasized and some of the claims are not fully supported by the data.

We have done our best to acknowledge other work on SUMO and its role in piRNA silencing. We apologize if it seems we were over-emphasizing our work at the expense of others. We have expanded the Discussion to improve the context of our work. The comment regarding our claims and data will be addressed below.

1. The importance of HDA-1 SUMOylation for transcriptional repression. The title "HDAC1 SUMOylation promotes Argonaute directed transcriptional silencing in C. elegans" implies a central role of SUMOylation in piRNA-mediated transcriptional silencing. The Argonaute HRDE-1/WAGO-9 targets countless transposons as shown previously and also in this manuscript (Figure S3), and so do the HDA-1 degron and Ubc9 mutant, indicating that histone deacetylation and protein SUMOylation are essential processes in TE silencing. However, the HDA-1 SUMOylation mutant (KKRR) only slightly affects 6 TE families (Figure S3), indicating that SUMOylation of HDA-1 might not be a key mediator of this process.

We have revised the manuscript to make this point more clearly. Gene silencing by the piRNA pathway includes two stages, initation and maintenance. The Piwi Argonaute PRG-1 is required for initiation not maintenance. While this role for PRG-1 in intiation was pieced together from transgene studies, the idea is that for most transposons, PRG-1 is/was required only to initially silence the transposon. Afterward the WAGO system maintains silencing indefinitely over many generations. We now explain this important detail better in the Results section.

Furthermore, the authors write that "Our findings suggest how SUMOylation of HDAC1 promotes the recruitment and assembly of an Argonaute-guided chromatin remodeling complex to orchestrate de novo gene silencing in the C. elegans germline.", but then they also state that "Comparison with mRNA sequencing data from auxin-treated degron::hda-1 animals revealed an even more extensive overlap with Piwi pathway mutants (Figure S2B), indicating that HDA-1 also promotes target silencing independently of HDA-1 SUMOylation."

We apologize for this confusing error. We sometimes think of the WAGO pathway as being a downstream part of the Piwi pathway, but to do so is confusing in this context. We revised this section to clarify the issue as follows:

“The silencing defect was more severe in auxin-treated degron::hda-1 than in hda-1[KKRR] (Figure 5—figure supplement 1B and Figure 5—figure supplement 2), resulting in the increased expression of many more transposons and a more extensive overlap with genes upregulated in rde-3 mutant worms (Figure 5—figure supplement 1C and Figure 5—figure supplement 2). This result indicates that HDA-1 also promotes the maintenance of silencing independently of HDA-1 SUMOylation.”

Based on their results and their own interpretations, I find that the importance of HDA-1 SUMOylation in piRNA-dependent transcriptional silencing is overemphasized.

Thanks for your very candid comments. I hope the above corrections help you understand the assay used here. The similarities between prg-1 and hda-1(KKRR) from our mRNA-seq data are very striking.

Additionally, the model (Figure 7) implies that for initiation of silencing WAGO recruits HDA-1 to targets. This should be tested by analyzing HDA-1 distribution over WAGO targets in WT and upon loss of WAGO.

As we hope is now clear, our model is that HDA-1 is transiently SUMOylated during the initial phase of converting an active heavily acetylated target gene into a silent state. The distribution of HDA-1, most of which would not be SUMOylated in any case, would be unlikely to provide any new insight.

2. The mechanistic role of HDA-1 SUMOylation.

On page 17 (amongst other places) the authors claim that "The SUMOylation of HDA-1 promotes its activity, while also promoting physical interactions with other components of a germline nucleosome-remodeling histone deacetylase (NuRD) complex, as well as the nuclear Argonaute HRDE-1/WAGO-9 and the heterochromatin protein HPL-2 (HP1)".

• Regarding activity: Loss of deacetylation/silencing in the SUMO mutant might be due to loss of enzymatic activity, but it might also be due to defects in recruitment/complex formation. There is no data that proves altered enzymatic activity.

We apologize again for the poor word choice. As stated above, we completely agree, and we did not mean to imply that SUMOylation regulates HDA-1 enzymatic activity. We have corrected this mistake throughout both papers.

In fact, Figure 6 indicates SUMO-dependent interaction of WAGO-9 with HDA-1, implying that recruitment is affected. To distinguish between activity and recruitment, at the very least, the authors would need to show that HDA-1 localization to its genomic targets is unaltered upon mutating its SUMOylation site (ChIP-seq of wt and KKRR mutant), while H3K9ac is increased (K9ac ChIP-seq in wt and KKRR mutant) in the mutant. This, in combination with HDA-1 localization in wt and WAGO-9 loss would imply whether complex formation to recruit HDA-1 or HDA-1 enzymatic activity is mostly affected by SUMOylation.

We hope it is now completely clear that we are not proposing that the SUMO modification alters enzyme activity. As detailed above, we have extensively revised the Discussion to address the above issues.

• Regarding physical interactions: Figure 3D shows that if we fuse a SUMO residue to HDA-1, it will interact with MEP-1, while SUMOylation deficient HDA-1 mutant doesn't interact. However, for the WT HDA-1 control, we only see unSUMOylated protein interacting with MEP-1. Furthermore, in the MEP-1 IPs of samples that should contain SUMO-fused HDA-1, the authors detect a lot of "cleaved", unSUMOylated HDA-1. Unless cleavage happened after IP, during elution (unlikely, and there is "cleaved" HDA-1 in the inputs), these findings argue that the interaction with MEP-1 is not mediated by HDA-1 SUMOylation. An interaction between MEP-1 and unmodified HDA-1 is also shown in the accompanying manuscript, which appears to be dependent on Pie-1 SUMOylation. Thus, SUMOylation of HDA-1 alone seems unlikely to be the major factor necessary for silencing complex assembly. (as a side question: Does the protease inhibitor cocktail used inhibit de-SUMOylation enzymes? I am concerned that deSUMOylating enzymes might compromise some result interpretations.)

Thanks again for your very critical read of our findings. As noted in the cover letter, we have dealt with the fleeting nature of this modification for so many years that—while developing this story—we forgot that it was not generally known or appreciated. In both studies, the SUMO modifications are completely removed when we prepare samples in IP buffer (even 5 minutes at 4°C). We only detect SUMO-conjugated proteins under stringent denaturing conditions. We now make a major discussion point of this interesting and perhaps underappreciated aspect of SUMO biology.

• Regarding functional relevance of HDA-1 acetylation: On pages 12/13 authors claim that because "HDA-1(KKRR) animals and mep-1-depleted worms revealed dramatically higher levels of H3K9Ac compared to wild-type" and "HDA-1, LET-418/Mi-2, and MEP-1 bind heterochromatic", "SUMOylation of HDA-1 appears to drive formation or maintenance of germline heterochromatin regions of the genome." These correlations do not prove function. The authors have performed H3K9me2 (although not H3K9-ac) ChIP-seq in WT, KKRR mutant and HDA-1 degron worms, yet do not analyze globally whether acetylation is lost on genes that are affected (change in RNA-seq vs. change in K9me2 or acetyl). To support the claim that SUMOylation of HDA-1 drives deacetylation and heterochromatin formation, it would be important to show changes in H3K9Ac levels (or other acetyl marks) and potentially NuRD component occupancy between control and HDA-1 SUMOylation-deficient animals at specific targets (i.e. genes derepressed upon loss of SUMOylation identified in RNA-seq, and the reporter locus).

As we note above, obtaining sufficient material for ChIP-seq studies on adult germline is technically difficult to do.

3. The authors claim (p17) that "initiation of transcriptional silencing requires SUMOylation of conserved C-terminal lysine residues in the type-1 histone deacetylase HDA-1". I do not see any supporting data that has separately looked at formation/initiation and maintenance of silencing (a technically challenging experiment).

Thanks again for pointing this out. The piRNA pathway initiates transcriptional silencing on actively transcribing genes, and it is in this context where HDA-1 SUMOylation appears to be most critical. We hope this issue is now explained better in the manuscript.

4. The authors repeatedly claim that gei-17 does not play a role in piRNA target silencing, based on loss of gei-17 not affecting the piRNA reporter (Figure 1B). At the same time, they claim that pie-1 plays a role, even though it likewise does not affect the piRNA reporter (it affects the reporter only in F3; data on gei-17 effect in F3 is not present). In the accompanying paper, the authors show that while gei-17 loss by itself causes only moderate effect on extra intestine cells, combined with Pie-1 loss the effect is more severe than when Pie-1 loss is combined with Ubc9 or smo loss. This to me indicates an important role of gei-17 in inhibiting differentiation of germline stem cells to somatic tissues, but these effects are likely synergistic and thus masked by Pie-1. Individually neither Gei-17 nor Pie-1 show an effect on piRNA reporter in P0, but to confirm lack of synergy, their effects of should be tested together. Although possible, the present data is insufficient to rule of gei-17 involvement.

You are correct. We have now completed double mutant studies that show that gei-17 and pie-1 function together to silence the piRNA sensor. This new data is in presented the PIE-1 paper, and discussed in both papers.

https://doi.org/10.7554/eLife.63299.sa2

Article and author information

Author details

  1. Heesun Kim

    RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Yue-He Ding
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7643-4516
  2. Yue-He Ding

    RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Heesun Kim
    Competing interests
    No competing interests declared
  3. Gangming Zhang

    RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, United States
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  4. Yong-Hong Yan

    National Institute of Biological Sciences, Beijing, China
    Contribution
    Investigation
    Competing interests
    No competing interests declared
  5. Darryl Conte Jr

    RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, United States
    Contribution
    Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1137-8901
  6. Meng-Qiu Dong

    National Institute of Biological Sciences, Beijing, China
    Contribution
    Supervision, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6094-1182
  7. Craig C Mello

    1. RNA Therapeutics Institute, University of Massachusetts Medical School, Worcester, United States
    2. Howard Hughes Medical Institute, Chevy Chase, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Investigation, Writing - original draft, Writing - review and editing
    For correspondence
    Craig.Mello@umassmed.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9176-6551

Funding

NIH Office of the Director (GM58800)

  • Craig C Mello

Howard Hughes Medical Institute

  • Craig C Mello

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

Acknowledgements

We thank members of the Mello and Ambros labs for discussions. We thank M Shirayama for sharing strains, and the Miska and Hay labs for sharing antibodies. Onur Yukselen and Alper Kucukural from Bioinformatics Core at UMass Medical School provided support on ChIP-seq data analysis. CCM is a Howard Hughes Medical Institute Investigator. This work was supported in part by NIH Grant GM 58800.

Senior Editor

  1. Kevin Struhl, Harvard Medical School, United States

Reviewing Editor

  1. Tony Hunter, Salk Institute for Biological Studies, United States

Reviewers

  1. Tony Hunter, Salk Institute for Biological Studies, United States
  2. Federico Pelisch, University of Dundee, United Kingdom

Publication history

  1. Received: September 21, 2020
  2. Accepted: April 23, 2021
  3. Version of Record published: May 18, 2021 (version 1)

Copyright

© 2021, Kim et al.

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

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