New Results

pals-22, a member of an expanded C. elegans gene family, controls silencing of repetitive DNA

Eduardo Leyva-Díaz, Nikolaos Stefanakis, Inés Carrera, Lori Glenwinkel, Guoqiang Wang, Monica Driscoll, Oliver Hobert
doi: https://doi.org/10.1101/145094

ABSTRACT

Repetitive DNA sequences are subject to gene silencing in various animal species. Under specific circumstances repetitive DNA sequences can escape such silencing. For example, when exogenously added, extrachromosomal DNA sequences that are stably inherited in multicopy repetitive arrays in the nematode C. elegans are frequently silenced in the germline, whereas such silencing often does not occur in the soma. This indicates that somatic cells might utilize factors that prevent repetitive DNA silencing. Indeed, such “anti-silencing” factors have been revealed through genetic screens that identified mutant loci in which repetitive transgenic arrays are aberrantly silenced in the soma. We describe here a novel locus, pals-22 (for protein containing ALS2CR12 domain), required to prevent silencing of repetitive transgenes in neurons and other somatic tissue types. pals-22 deficiency also severely impacts animal vigor and confers phenotypes reminiscent of accelerated aging. We find that pals-22 is a member of a large family of divergent genes (39 members), defined by the presence of an ALS2CR12 domain. While gene family members are highly divergent, they show striking patterns of genomic clustering. The family expansion appears C. elegans-specific and has not occurred to the same extent in other nematode species. Previous transcriptome analysis has revealed that most of the pals genes are induced under stress conditions or upon infection by intracellular parasites. The transgene silencing phenotype observed upon loss of cytoplasmically localized PALS-22 protein depends on the biogenesis of small RNAs, since silencing is abolished in the RNAi defective mutant rde-4, suggesting that pals-22 might regulate RNAi dependent silencing in the cytoplasm of neurons and other tissues. We speculate that the pals gene family may be part of a species-specific cellular defense mechanism.

INTRODUCTION

Over half the human genome consists of repetitive DNA elements (Landeret al. 2001; de Koninget al. 2011). The view of the role of repetitive DNA has evolved in the last decades from considering it as “junk DNA” to the recognition of repetitive DNA as essential for genome function (Doolittle AND Sapienza 1980; Orgel AND Crick 1980; Lynch AND Conery 2003; Shapiro AND von Sternberg 2005). The main constituents of these repetitive DNA elements are retrotransposons, a large family of transposable elements capable of copying themselves and reinserting into the host genome (Kazazian 2004; Goodier AND Kazazian 2008; Cordaux AND Batzer 2009). Retrotransposons and other elements with the ability to copy themselves pose a threat to genome integrity due to the potential deleterious effects of landing in coding or regulatory regions (Friedli AND Trono 2015). The activation of proto-oncogenes in some leukemias represent an example of such harmful consequences (Hacein-Bey-Abinaet al. 2003). However, repetitive DNA elements have also been found to play beneficial roles in a number of processes, ranging from the regulation of gene expression to interaction with nuclear structures for genome packaging, to DNA repair and restructuring (Shapiro AND von Sternberg 2005; Goke AND Ng 2016). Moreover, retrotransposons are a source of regulatory elements and alternative gene isoforms. Hence, transposable elements have been postulated as a powerful genetic force involved in the evolution of organismal complexity (Kazazian 2004; Friedli AND Trono 2015).

To balance the deleterious and beneficial features of repetitive sequences, organisms have evolved ways to finely tune the regulation of repetitive DNA elements (Schlesinger AND Goff 2015; Chuonget al. 2017). For example, endogenous silencing mechanisms have evolved to prevent genome damage by the spread of mobile repetitive DNA elements. Silencing is achieved by DNA binding proteins, histone modifications and DNA methylation, both of which can be directed by sequence-specific repressive transcription factors and by small RNAs. For example, retrotransposons are extensively recognized by the Krueppel-associated box-zinc finger (KRAB-ZFP) proteins (Roweet al. 2010; Quennevilleet al. 2012; Jacobset al. 2014), which form a large family of repressive transcription factors. These transcription factors are among the fastest evolving group of genes in the human genome and their diversity facilitates their ability to recognize a large number of retrotransposons (Nowicket al. 2010). Sequence-specific binding to retrotransposons by KRAB-ZFP factors triggers a cascade leading to chromatin-based silencing mechanisms (Wolf AND Goff 2009).

RNAs of retrotransposons that escape transcriptional silencing are targeted and destroyed by the small RNA pathways in the cytoplasm (Tothet al. 2016). RNA-based mechanisms represent the most ancient defense against the genomic spread of repetitive DNA elements (Friedli AND Trono 2015). These mechanisms comprise the action of small RNA molecules, including small interfering RNAs (siRNAs), PIWI-interacting RNAs (piRNAs) and microRNAs (miRNAs), which guide repressor protein complexes to particular targets in a sequence-specific manner. In addition to intervening at the post-transcriptional level, small RNAs can also intervene at the transcriptional level by directing deposition of repressive histone marks and DNA methylation to copies of retrotransposons and other elements (Le Thomaset al. 2013). piRNAs and siRNAs can be produced from repetitive DNA elements, which they silence in return (Law AND Jacobsen 2010). Thus, in many organisms, repetitive DNA serves as a trigger for gene silencing.

Repetitive DNA elements are not only abundant in vertebrate genomes. Repetitive DNA elements also abound in model organisms with comparatively smaller genome sizes: repetitive DNA accounts for 34-57% of the total genome in Drosophila melanogaster (Celnikeret al. 2002), and at least 17% of the Caenorhabditis elegans genome (Steinet al. 2003). Repetitive DNA elements can also be generated experimentally. DNA transformation techniques in C. elegans produce repetitive extrachromosomal DNA arrays (“simple” arrays)(Melloet al. 1991). Several studies have shown that expression of transgenes organized in these repetitive arrays is silenced both in somatic cells and in the germline through heterochromatin formation, involving several chromatin factors (reviewed in (Cui AND Han 2007)). Somatic, and especially, germline transgene expression can be improved when the transgenic DNAs are cotransformed with an excess of carrier DNA, producing a less repetitive, more “complex” array (Kellyet al. 1997).

Importantly, gene expression from repetitive genomic regions can still be observed, suggesting there are mechanisms that can modulate silencing effects (Tsenget al. 2007). Multiple forward genetic screens in C. elegans have indeed identified factors that act to counter silencing of genes contained in repetitive sequences, based on screens for mutations that alter the activity of transgenes present in tandemly repeated arrays. (Hsiehet al. 1999; Grishoket al. 2005; Tsenget al. 2007; Fischer et al. 2013). In a classic study, mutations in tam-1 (a RING finger/B-box factor) were found to reduce the expression of transgenes organized in simple but not complex repetitive arrays (Hsiehet al. 1999). Therefore, tam-1 is an “anti-silencing factor” that attenuates the context-dependent silencing mechanism affecting multi-copy tandem-array transgenes in C. elegans. A subsequent study identified mutations in another gene important for expression of repetitive sequences, lex-1, which genetically interacts with tam-1 (Tsenget al. 2007). LEX-1 encodes a protein containing an ATPase domain and a bromodomain, both of which suggest that LEX-1 associates with acetylated histones and modulates chromatin structure. Hence, TAM-1 and LEX-1 are anti-silencing factors that function together to influence chromatin structure and to promote expression from repetitive sequences. Further studies found that tandem-array transgenes become silenced in most mutants that cause enhanced exogenous RNAi (Simmeret al. 2002; Kennedyet al. 2004; Fischeret al. 2013). Examples of gene inactivations known to cause increased transgene silencing and enhanced RNAi include the retinoblastoma like gene lin-35 (Hsiehet al. 1999; Wanget al. 2005; Lehneret al. 2006), the RNA-dependent RNA polymerase rrf-3 (Simmeret al. 2002), and the helicase gene eri-6/7 (Fischeret al. 2008). Silencing of repetitive DNA elements (multi-copy transgenes) depends on a complex interaction between different small RNA pathways (Fischeret al. 2013).

Here, we identify a novel locus, pals-22, whose loss confers a transgene silencing phenotype. pals-22 mutants display context-dependent array silencing, affecting the expression of highly repetitive transgenes but not single copy reporters. Animals lacking pals-22 show locomotory defects and premature aging. pals-22 is a member of a large family of divergent genes defined by the presence of an ALS2CR12 domain. The ALS2CR12 domain protein family is specifically expanded in C. elegans, and pals gene family members are clustered in the genome. We found that transgene silencing on pals-22 mutants depends on the RNAi pathway, indicating that pals-22 might act as regulator of small RNA-dependent gene silencing.

MATERIALS AND METHODS

Mutant strains

Strains were maintained by standard methods (Brenner 1974). The C. elegans mutant alleles used in this study were: pals-22(ot723), pals-22(ot810), pals-22(ot811), rde-4(ne301) (Tabaraet al. 1999) and tam-1(cc567) (Hsiehet al. 1999).

Reporter and transgenic strains

The C. elegans transgenic strains used in this study were otls381[ric-19prom6::NLS::gfp], otls380[ric-19prom6::NLS::gfp], ccls4251[myo-3prom::gfp], otls251[cat-2prom::gfp], otIs355[rab-3prom1::NLS::rfp], otls447[unc-3prom::mChOpti], otEx6944 [ric-4prom26::NLS::yfp], otls620[unc-11prom8::NLS::gfp], otls353[ric-4fosmid::SL2::NLS-YFP-H2B], otls534[cho-1fosmid::SL2::NLS-YFP-H2B], otTi32[lin-4prom::yfp], ieSi60[myo-2prom::TlR1::mRuby], otEx7036[pals-22prom::gfp], otEx7037 [pals-22::gfp]. pals-22 GFP reporters were generated using a PCR fusion approach (Hobert 2002). Genomic fragments were fused to the GFP coding sequence, which was followed by the unc-54 3′ UTR. See Table 1 for transgenic strain names and microinjection details.

View this table:
Table 1:

Transgenic strains used in this study.

Forward genetic screens

Standard ethyl methanesulfonate (EMS) mutagenesis was performed on the fluorescent transgenic reporter strain: otls381[ric-19prom6::NLS::gfp] and ~60,000 haploid genomes were screened for expression defects with an automated screening procedure (Doitsidouet al. 2008) using the Union Biometrica COPAS FP-250 system. To identify the causal genes of the mutants obtained, we performed Hawaiian single nucleotide polymorphism (SNP) mapping and whole-genome sequencing (Doitsidouet al. 2010) followed by data analysis using the CloudMap pipeline (Minevichet al. 2012). The following mutant alleles were identified: pals-22(ot810) and pals-22(ot811). The mutant allele pals-22(ot723) was identified in an independent manual clonal screen for changes in reporter expression in neurons of otls381[ric-19prom6::NLS::gfp] after EMS mutagenesis.

Microscopy

Worms were anesthetized using 100 mM sodium azide (NaN3) and mounted on 5% agarose pads on glass slides. All images (except Figure 3, B-C, and Figure S1C) were acquired as Z-stacks of ~1 μm-thick slices with the Micro-Manager software (Edelsteinet al. 2010) using the Zeiss Axio Imager.Z1 automated fluorescence microscope. Images were reconstructed via maximum intensity Z-projection of 2-10 μm Z-stacks using the ImageJ software (Schneideret al. 2012). Images shown in Figure 3, B-C, and Figure S1C were acquired using a Zeiss confocal microscope (LSM880). Several z-stack images (each ~ 0.4 µm thick) were acquired with the ZEN software. Representative images are shown following orthogonal projection of 2-10 µm z-stacks.

Single molecule FISH

smFISH was done as previously described (Ji AND van Oudenaarden 2012). Samples were incubated overnight at 37°C during the hybridization step. The ric-19 and gfp probes were designed using the Stellaris RNA FISH probe designer and was obtained conjugated to Quasar 670 from Biosearch Technologies.

Fluorescence quantification

Synchronized day 1 adult worms were grown on NGM plates seeded with OP50 and incubated at 20°C. The COPAS FP-250 system (Union Biometrica) was used to measure the fluorescence of 200-1000 worms for each strain.

Bioinformatic analysis

The ALS2CR12 domain phylogenetic tree was generated using MrBayes (Huelsenbeck AND Ronquist 2001; Ronquist AND Huelsenbeck 2003), lset nst=6 rates=invgamma, ngen increased until the standard deviation of split frequencies < 0.05. Input protein coding sequences for ALS2CR12 domain and PALS family orthologous proteins were aligned with M-Coffee (Wallaceet al. 2006). The MrBayes tree figure was rendered with FigTree (http://tree.bio.ed.ac.uk/software/figtree/).

RNAi by feeding

RNAi was performed as previously described with minor adaptations (Kamath AND Ahringer 2003). L4-stage hermaphrodite worms were placed onto NGM plates containing seeded bacteria expressing dsRNA for each assayed gene. After 24 h at 20°C, adults were removed. After a further 36-40 h at 20°C, phenotypes were scored blindly.

Computer swim analysis

The swimming assay was performed using the CeleST program as previously described (Restifet al. 2014). In brief, we transferred five (day-1) adult hermaphrodites into 50 μl M9 buffer located in a 10 mm staggered ring on a glass slide. A 30 second dark-field video (18 frames per second) was immediately recorded via StreamPix 7. Multiple features of the swim behavior were then analyzed using CeleST (Restifet al. 2014). Graphpad Prism 6 was used for data plotting and statistics.

Crawling assay

We washed ~30 day-1 adult hermaphrodites into M9 buffer (containing 0.2% BSA to prevent worms sticking to plastic tips) via low speed centrifuging. We transferred these worms (in 20 μl volume) to a NGM agar plate (60 mm). After the liquid was completely dried and most animals were separated from each other, we started a 30 second video recording (20 frames per second). The video was processed on ImageJ and analyzed via wrMTrck plugin (Nussbaum-Krammeret al. 2015). The crawling paths were generated in ImageJ and enhanced with Photoshop.

Age pigment assay

Age pigments of day-5 adult hermaphrodite (Gerstbreinet al. 2005) were captured via Zeiss LSM510 Meta Confocal Laser Scanning Microscope (excitation: Water cooled Argon laser at 364nm; emission: 380nm–420nm). The auto-fluorescence intensity was quantified in ImageJ.

Lifespan

Synchronized worms were picked at the L4 stage, and fed with OP50-1 bacteria on a 35mm NGM agar plate (12 worms per plate, ~100 animals initiating each trial). Before the end of reproductive phase, animals were transferred into a new plate every two days to keep adults separated from progeny. Immobile animals without any response to touch were counted as dead; bagged worms were also counted as deaths; animals crawling off the NGM agar were counted as lost and were excluded from analysis.

RESULTS

pals-22 mutants show a transgene silencing phenotype

Based on our long-standing interest in studying the regulation of pan-neuronal gene expression (Stefanakiset al. 2015), we sought to use genetic mutant screens to isolate factors that control the expression of pan-neuronally expressed reporter transgenes. One screen that we undertook used a regulatory element from the pan-neuronally expressed ric-19 locus, fused to gfp (otIs381[ric-19prom6::NLS::gfp])(Stefanakiset al. 2015). We identified three independent mutant alleles, ot723, ot810 and ot811, in which expression of ric-19prom6::NLS::gfp was reduced throughout the nervous system in all animals examined (Fig. 1A, B). Using our previously described whole-genome sequencing and mapping pipeline (Doitsidouet al. 2010; Minevichet al. 2012), we found that all three mutations affect the same locus, C29F9.1 (Fig. 2A, B), which we named pals-22 for reasons that we explain further below. The ric-19prom6::NLS::gfp expression defect of pals-22(ot811) can be rescued by a fosmid (WRM0616DC09) encompassing the pals-22 locus plus neighboring genes as well as a genomic fragment that only contains the pals-22 locus (791 bp upstream of the start codon to the stop codon and its 3’UTR)(Fig. 2C). Both ot810 and ot811 alleles carry early nonsense mutations and are therefore predicted to be null alleles (Fig. 2B).

Fig. 1:
Fig. 1: Loss of pan-neuronal reporter gene expression in pals-22 mutants.

(A) The ric-19prom6::NLS::gfp transcriptional reporter is brightly expressed in all neurons in wild-type N2 worms, with a nuclear localization. In pals-22(ot723), pals-22(ot810) and pals-22(ot811) mutants, expression is reduced throughout all the nervous system. All images correspond to L4 worms. Scale bar represents 50 μm

(B) Fluorescence profiles of a representative individual day 1 adult worm expressing ric-19prom6::NLS::gfp in wild-type (black line) or pals-22(ot811) mutant background (red line) obtained with a COPAS FP-250 system. Time of flight indicates worm length, with lower values corresponding to the head of the worms. a.u., arbitrary units. Inset bar graph displays quantification of total fluorescence intensity averaged over 500 animals analyzed by COPAS. The data are presented as mean + SEM. Unpaired t-tests were performed for pals-22(ot811) compared to WT; ***p < 0.001.

(C) Wild-type (left), pals-22(ot810) (middle) and pals-22(ot811) (right) images of the anterior part of L3 worms showing equal ric-19 mRNA levels in control and mutant worms as assessed by single molecule fluorescence in situ hybridization. Individual transcripts shown as purple dots in top and as black dots in bottom panels. GFP expression of the reporter transgene is shown in green. At least 20 animals examined for each genotype displayed indistinguishable staining. Scale bar 10 μm.

Fig. 2:
Fig. 2: pals-22 codes for a protein with an ALS2CR12 domain.

(A) Hawaiian single nucleotide polymorphism (SNP) mapping plots obtained from whole-genome sequencing of the following mutant alleles: pals-22(ot810) (top row), pals-22(ot811) (middle row), and pals-22(ot723) (bottom row).

(B) Schematic of pals-22 gene locus depicting its ALS2CR12 domain and mutant allele annotation.

(C) pals-22 rescue data. %WT = % percent animals that express the ric-19 reporter strongly (as in wild-type animals). %MUT = % percent animals that express the ric-19 reporter more weakly than wild-type.1otIs381 = ric-19prom6::NLS::gfp.2WRM0616DC09 fosmid.

pals-22 mutants display reduced GFP expression of two separate ric-19prom6::NLS::gfp integrated reporter transgenes (Table 2). However, single molecule fluorescence in situ hybridization (smFISH) against endogenous ric-19 transcripts failed to detect effects on the endogenous ric-19 expression (Fig. 1C). Since the two ric-19 reporter transgenes that are affected by pals-22 are repetitive, “simple” arrays, we considered the possibility that pals-22 may encode a transgene silencing activity. To test this notion, we examined the expression of a wide range of reporter transgenes in pals-22-deficient mutants (summarized in Table 2). Six additional simple arrays with widely different cellular specificities of expression (pan-neuronal, dopaminergic neurons, ventral cord motorneurons, muscle) are also silenced in pals-22 mutants (Table 2, Fig. 3). Two of these arrays, myo-3::gfp (ccIs4251), and cat-2::gfp (otIs251) were previously shown to be silenced by loss of tam-1, a “classic” transgene silencer mutation (Hsiehet al. 1999)(M. D. and O. H. unpublished data)(Fig. 3). We quantified the magnitude of the pals-22(ot811) effect on expression of simple array reporters by acquiring fluorescence intensity information from a synchronized worm population of worms using a COPAS FP-250 system (Union Biometrica; “worm sorter”). At the L4 larval stage, we observed a 76% reduction in green fluorescence intensity for ric-19prom6::NLS::gfp, 66% reduction for myo-3::gfp, 32% reduction for cat-2::gfp and 42% reduction in red fluorescence intensity for rab-3prom1::NLS::rfp (Fig. 1B; Fig. 3B, D, E).

Fig. 3:
Fig. 3: pals-22 mutants show silencing of several multicopy arrays.

(A) The myo-3prom::gfp transcriptional reporter is brightly expressed in all body muscles in wild-type N2 worms, with a combination of mitochondrial and nuclear localization. In pals-22(ot810), pals-22(ot811) and tam-1(cc567) mutants a generalized reduction in GFP fluorescence is observed. All images correspond to L4 worms. Scale bar represents 50 μm.

(B) Fluorescence profile of a representative individual day 1 adult worm expressing myo-3prom::gfp in wild-type (black line) or pals-22(ot811) mutant background (red line) obtained with a COPAS FP-250 system. Time of flight indicates worm length, with lower values corresponding to the head of the worms. a.u., arbitrary units. Inset bar graph displays quantification of total fluorescence intensity averaged over 1000 animals analyzed by COPAS. The data are presented as mean + SEM. Unpaired t-tests were performed for pals-22(ot811) compared to WT; ***p < 0.001.

(C) GFP images showing silencing of cat-2prom::gfp expression in the head of pals-22(ot811) mutants (right) compared to wild-type L4 worms (left). The cat-2prom::gfp transcriptional reporter is expressed in all dopamine neurons, CEPD, CEPV and ADE in the head, and PDE in the posterior mid-body (top right insets). Scale bar represents 10 μm.

(D) Fluorescence profile of a representative individual day 1 adult worms expressing cat-2prom::gfp in wild-type (black line) or pals-22(ot811) mutant background (red line) obtained with a COPAS FP-250 system. Time of flight indicates worm length, with lower values corresponding to the head of the worms. a.u., arbitrary units. Inset bar graph displays quantification of total fluorescence intensity averaged over 1000 animals analyzed by COPAS. The data are presented as mean + SEM. Unpaired t-tests were performed for pals-22(ot811) compared to WT; ***p < 0.001.

(E) RFP images showing silencing of rab-3prom1::rfp expression in the head of pals-22(ot811) mutants (right) compared to wild-type L4 worms (left).

(F) Fluorescence profile of a representative individual day 1 adult worms expressing rab-3prom1::rfp in wild-type (black line) or pals-22(ot811) mutant background (red line) obtained with a COPAS FP-250 system. Time of flight indicates worm length, with lower values corresponding to the head of the worms. a.u., arbitrary units. Inset bar graph displays quantification of total fluorescence intensity averaged over 1000 animals analyzed by COPAS. The data are presented as mean + SEM. Unpaired t-tests were performed for pals-22(ot811) compared to WT; ***p < 0.001.

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Table 2:

Pals-22 effects on transgene reporter expression.

We also analyzed the expression of complex array (tandemly repeated transgenes with a less repetitive structure) and single copy reporter transgenes in pals-22 mutants (summarized in Table 2). One complex array transgene was silenced (unc-11p8::gfp, Table 2) but others were not (ric-4fosmid::yfp and cho-1fosmid::yfp, Table 2, Fig. S1A). Perhaps the much smaller reporter fragment unc-11p8::gfp generated more repetitive structures even in the context of “complex” arrays than the larger fosmid-based reporters. Importantly, single copy insertions or endogenously tagged genes are not affected by mutations in pals-22 (Fig. S1B, C).

Similar to previously characterized transgene silencing mutants, such as tam-1 (Hsiehet al. 1999), the reduction of reporter expression is temperature sensitive. However, the direction of the sensitivity is inverted as compared to the tam-1 case: the decrease in ric-19prom6::NLS::gfp expression is most pronounced at 15°C, while the effect is milder at 25°C (data not shown). We also found stage dependent variability: the decrease in transgene reporter expression is stronger as the animals develop, from mild differences in expression in early stages of development to more obvious defects at later stages (Fig. S2).

PALS-22 is a broadly expressed, cytoplasmic protein

To analyze the expression pattern of pals-22 we fused the entire locus to gfp (including 791 bp of 5’ sequences and all exons and introns; Fig. 4A). This reporter construct fully rescues the transgene silencing phenotype of pals-22(ot811) mutants (Fig. 4D). Expression is observed widely across different tissues: nervous system (pan-neuronal expression), body wall and pharyngeal muscle, gut, seam cells, as well as the male tail (Fig. 4B, C). A similar expression pattern is observed with a transcriptional reporter (791 bp of its upstream region fused to GFP; Fig. 4C). The rescuing, translational reporter transgene revealed a strong, if not exclusive enrichment in the cytoplasm and appears to be excluded from the nucleus in most tissues (Fig. 4B). However, we cannot exclude the possibility that the nucleus contains low amount of functional PALS-22 protein.

Fig. 4:
Fig. 4: PALS-22 is a broadly expressed, cytoplasmic protein.

(A) Schematic of pals-22 transcriptional and translational GFP reporters. The gfp reporter is followed by the 3’UTR from the unc-54 gene.

(B) Expression of the pals-22::gfp translational reporter (otEx7037) is shown in green in the head (left) in a representative hermaphrodite L4 worm. High magnification images in black and white for the head (middle) and tail (right) show PALS-22 cytoplasmic localization. Nuclear pan-neuronal rab-3prom1::rfp expression is shown in red on the background. Two distinct extrachromosomal arrays show the same pattern.

(C) Expression of the pals-22prom::gfp transcriptional reporter (otEx7036) is shown in green in the head (left panel), mid-body (second panel), and tail (third panel) in a representative hermaphrodite L4 worm; and in the male tail (right panel). Nuclear pan-neuronal rab-3prom1::rfp expression is shown in red on the background. Three distinct extrachromosomal arrays show the same pattern.

(D) PALS-22::GFP (right) rescues the expression of unc-3prom::mChOpti in pals-22(ot811) mutants (left). At least 50 animals examined for each genotype. Scale bars represent 20 μm (B and C), 5 μm (B, high magnification images) and 50 μm (D).

pals-22 is a member of an unusual C. elegans gene family

The molecular analysis of the PALS-22 protein sequence revealed its membership in an unusual gene family. The only recognizable feature of PALS-22 is a domain termed ALS2CR12 by the InterPro (https://www.ebi.ac.uk/interpro/) (Finnet al. 2017) and Panther (http://www.pantherdb.org/)(Miet al. 2013; Miet al. 2017) databases (hence the name “PALS” for “protein containing ALS2CR12 domain”). This domain was first found in a human gene that constituted a potential disease locus for amyotrophic lateral sclerosis 2 (Hadanoet al. 2001). No biochemical or cellular function has yet been assigned to the human ALS2C12 protein or any of its homologs in other organisms.

Both InterPro and Panther databases group PALS-22 within a family of ALS2CR12 domain containing proteins from different species ranging from nematodes to vertebrates (Fig. 5A). While there is only one ALS2CR12 domain containing protein in vertebrates such as mouse or human, the number of proteins containing this domain is strikingly expanded to a total of 39 distinct proteins in C. elegans (Fig. 5A; Table 3). Drosophila seems to be completely devoid of ALS2CR12 domain-containing proteins.

Fig. 5:
Fig. 5: Sequence and genomic analysis of pals gene family in C. elegans and other species.

(A) Number of genes containing the ALS2CR12 domain in their protein product as predicted by InterPro (IPR026674) and/or Panther (PTHR21707).

(B) Graph representing the C. elegans to C. briggsae domain frequency for domains enriched in C. elegans (blue), C. briggsae to C. elegans ratio for domains enriched in C. briggsae (orange), or white for Panther protein domains predicted in the same number of genes in both species. Boxes indicate the gene counts for highly enriched domains in C. elegans. The number of Panther domain hits for the C. elegans and C. briggsae genome were obtained from WormBase.

(C) Phylogram of ALS2CR12 domain containing genes, including C. elegans paralogs and orthologs from C. briggsae, C. remanei, C. brenneri, human and mouse. Node values indicate posterior probabilities for each split as percent. The scale bar indicates average branch length measured in expected substitutions per site.

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Table 3:

List of all pals genes

The 39 C. elegans PALS proteins are very divergent from one another, as reflected in Table 3 with similarity scores of PALS-22 compared to other PALS proteins. Besides poor paralogy, there is also poor orthology. For example, BLAST searches with PALS-22 picks up no sequence ortholog in C. briggsae, and the best homolog from another species (C. brenneri, CBN22612) is less similar to PALS-22 than some of the C. elegans PALS-22 paralogs. The divergence of C. elegans pals genes is also illustrated by the genome sequence of wild isolates of C. elegans in which many pals genes display an above-average accumulation of polymorphisms (Volkerset al. 2013)(Table 3).

The expansion of C. elegans ALS2CR12 domain-containing proteins appears to be nematode species-specific, as C. briggsae only contains 8 predicted proteins with an ALS2CR12 domain; other nematodes also contain significantly less ALS2CR12 domain proteins (Fig. 5A). Such C. elegans-specific gene expansion is highly unusual, as shown in Fig. 5B. Among 3874 Panther protein domains analyzed, 2759 (71.2%) are present in the same number of genes in both C. elegans and C. briggsae, 128 (3.3%) domains are present only in C. elegans, while 74 (1.9%) are present only in C. briggsae. Most of the remaining domains are only slightly enriched in one species or the other. Just 10 domains (0.3%) are enriched five times or more in C. elegans versus C. briggsae (Fig. 5B). Most of these domain families contain uncharacterized genes, even though many of them contain human and vertebrate orthologs. Among them, the ALS2CR12 domain has not only a noteworthy enrichment, but it also constitutes the family with the largest absolute number of genes (Fig. 5B).

As perhaps expected from a species-specific expansion, the ALS2CR12 domainencoding pals genes are genomically clustered (Fig. 6). 14 genes are clustered in chromosome I (position I: 17.22 cM); while three clusters with 7 genes (III: -21.90 cM), 4 genes (III: -26.98 cM) and 3 genes (III: -3.18 cM) are present in chromosome III; lastly 4 genes are clustered in chromosome V (V: 4.22) (Fig. 6; Table 3). Taking into account the C. elegans-specific family expansion it is not surprising to find a total lack of conservation in the regions encompassing most of the pals clusters (Fig. 6). Only one cluster on chromosome V shows some degree of conservation among other nematode species. Genes surrounding these regions are conserved, suggesting recent gene duplications in the non-conserved areas. The low conservation region in cluster III: - 21.90 cM, contains many additional, non-conserved genes that are largely expanded in a nematode-specific manner, namely the previously analyzed fbxa genes (Thomas 2006).

Fig. 6:
Fig. 6: C. elegans pals genes are clustered and these clusters are poorly conserved.

Schematics of different C. elegans genomic regions adapted from the UCSC Genome Browser (https://genome.ucsc.edu/). For each panel is represented, from top to bottom:C elegans genome assembly (RefSeq genes) and conservation of 26 distinct nematode species (Basewise Conservation by PhyloP track). One isoform per gene is shown. pals genes are indicated in red, fbxa genes in blue. The following regions are shown:

Chromosome I, cluster at position 17.22 cM, chrl: 13,099,564-13,160,497 bp.

Chromosome Ill, cluster at position -21.90 cM, chrIII: 1,215,665-1,423,550 bp.

Chromosome III, cluster at position -26.98 cM, chrIII: 89,907-159,962 bp.

Chromosome III, cluster at position -3.18 cM, chrIII: 4,368,479-4,405,090 bp.

Chromosome V, cluster at position 4.22 cM, chrV: 12,427,096-12,462,015 bp.

Local gene duplications seem a plausible mechanism for the origin of the expanded C. elegans pals gene family. Consequently, we reasoned that pals genes within the same cluster should be more similar among each other than to other pals. In order to explore this possibility, we built a phylogenetic tree to visualize the phylogenetic relationships between pals genes (Fig. 5C). We included all C. elegans pals genes plus orthologs from C. briggsae, C. remanei, C. brenneri, mouse and human (based on presence of InterPro IPR026674). As expected, pals genes within the same cluster have a closer phylogenetic relationship, suggesting a shared origin.

According to modENCODE expression data (Celniker et al. 2009), most of the genes within each cluster are expressed in the same stage (e.g. all genes clustered in chromosome I are only found in L4 males), suggesting related functions (Table 3). Perhaps most intriguingly, though, the majority of pals genes become upregulated upon exposure to specific pathogens, specifically the exposure to intracellular fungal pathogen (microsporidia) or by viral infection (Bakowskiet al. 2014; Chenet al. 2017). Induction of pals gene expression is also observed upon various other environmental insults (exposure to toxic compounds)(Cuiet al. 2007)(summarized in Table 3). Several fbxa genes present in the low conservation region in cluster III: -21.90 cM, also become upregulated upon exposure to microsporidia or by viral infection (Bakowskiet al. 2014; Chenet al. 2017).

Somatic transgene silencing in pals-22 mutants requires rde-4-dependent small RNAs

We examined whether two pals-22 paralogs, pals-19 and pals-25 may also be involved in transgene silencing. Both genes show significant sequence similarity to pals-22 (Table 3) and one (pals-25) is directly adjacent to pals-22 (Fig.6). We tested whether two nonsense alleles generated by the Million Mutant project, pals-19(gk16606) and pals-25(gk891046) (Thompsonet al. 2013) silence the ric-19prom6::NLS::gfp and myo-3::gfp multi-copy transgenes. Neither array shows obvious changes in expression in pals-19 or pals-25 mutant backgrounds (data not shown).

To further pursue the role of pals-22 in transgene silencing we considered previous reports on transgene silencer mutations. Several genes with transgene silencing effects are known to be involved in modifying chromatin (Cui AND Han 2007). However, the cytoplasmic localization of PALS-22 described above argues against a direct role in controlling chromatin architecture (although, a function in the nucleus can not be ruled out). Nevertheless, we do find that pals-22 affects transgene silencing on the transcriptional level. smFISH against gfp mRNA shows that silenced gfp transgenic arrays display a significantly reduced number of transcripts in pals-22 mutants (Fig. 7A). We therefore considered the possibility that the transcriptional effects of a cytoplasmic protein on transcription may be controlled by intermediary factors. Small interfering RNAs are known to affect gene expression on the transcriptional level (Zamoreet al. 2000; Elbashiret al. 2001) and through a genetic epistasis test, we asked whether pals-22 requires small RNAs for its function. To this end, we turned to rde-4 mutant animals. The dsRNA-binding protein RDE-4 initiates gene silencing by recruiting an endonuclease to process long dsRNA into short dsRNA and is involved in exogenous as well as endogenous RNAi pathways (Tabaraet al. 2002; Parkeret al. 2006; Gentet al. 2010; Vasaleet al. 2010). We find that loss of rde-4 completely suppresses the pals-22 mutant phenotype (Fig. 7B, C). This result demonstrates that the gene silencing mediated by pals-22 deficiency requires the production of small dsRNAs.

Fig. 7:
Fig. 7: Somatic transgene silencing in pals-22 depends on the RNAi pathway.

(A) Wild-type (left), pals-22(ot810) (middle) and pals-22(ot811) (right) images of the anterior part of L3 worms showing reduced gfp mRNA levels in mutants compared to control wild-type worms as assessed by single molecule fluorescence in situ hybridization. lndividual transcripts shown as purple dots in top and as black dots in bottom panels. GFP expression is shown in green, DAPl staining is shown in blue. At least 20 animals examined for each genotype.

(B) Silencing of ric-19prom6::NLS::gfp in pals-22(ot811) (top) is suppressed in pals-22(ot811);rde-4(ne301) double mutants (bottom).

(C) Quantification of the data represented in (B). The data are presented as mean + SEM. Unpaired t-test, ***p < 0.001; n ≥ 300-1000 for all genotypes.

(D) Animals of the indicated genotype were grown on bacteria expressing dpy-13 or cel-1 dsRNA (feeding RNAi). For dpy-13 (top), progeny were scored for the percentage of animals having a dumpy body shape (Dpy). For cel-1 (bottom), the percentage of their progeny arresting at the L2 larval stage was determined. Wild-type otIs381[ric-19prom6::NLS::gfp] and rrf-3(pk1426) mutants were used as negative and positive controls. The data are presented as mean + SEM among the data collected from at least four independent experiments. Unpaired t-tests were performed for pals-22(ot811) and rrf-3(pk1426) compared to WT; ***p < 0.001, **p < 0.01, *p < 0.05.

Scale bars represent 10 μm (A) and 50 μm (B).

Transgene silencing phenotypes have been observed in mutants that affect multiple distinct small RNA pathways, and exogenous RNAi responses are often enhanced in these mutants (Simmeret al. 2002; Lehneret al. 2006; Fischeret al. 2013). Thus, we tested whether pals-22(ot811) shows an enhanced exogenous RNAi response. Using rrf-3(pk1426) as a positive control (Simmeret al. 2002), we detect enhanced dpy-13 or cel-1 RNAi phenotypes in pals-22(ot811) (dsRNA delivered by feeding; Fig. 7D). We conclude that pals-22 physiological function might be related to the regulation of RNAi-dependent silencing in the cytoplasm, via a mechanism critical to its action as an anti-silencing factor.

Locomotory and aging defects of pals-22 mutant animals

The loss of pals-22 has striking physiological consequences. Since casual observation of pals-22 mutants indicates defects in locomotion, we quantified these defects, by measured swimming behavior (Restifet al. 2014) and crawling activity, comparing wild-type, pals-22(ot810), pals-22(ot811) and pals-22(ot811) carrying a wild-type copy of pals-22 on an extrachromosomal array (Fig. 8). Day 1 adult pals-22 mutants show a poor performance in swimming assays as evaluated by multiple parameters, including low wave initiation rate (akin to thrash speed), travel speed (distance moved over time), brush stroke area (area covered in unit time) and activity index (Fig. 8A). On agar plates animals also were clearly impaired, displaying significantly decreased traveling speed (Fig. 8B). All locomotory defects were rescued by the pals-22(+) extrachromosomal array.

Fig. 8:
Fig. 8: pals-22 mutants show defective locomotion and early onset of aging traits.

(A) Day-1 adult pals-22 mutants exhibit defective swimming features, including decreased wave initiation rate, travel speed, brush stroke and activity index. pals-22(PCR_R) is the pals-22(ot811) mutant carrying a wild-type copy of pals-22 on an extrachromosomal array amplified by PCR. Data shown are mean ± SEM of each parameters, n = 30-35 (number indicated in each bar) from two independent experiments, *** p < 0.0001 (one-way ANOVA) compared to related control.

(B) pals-22 mutants crawl slowly on agar plates. The left panel shows the crawling path of each animal in 30 seconds. The right panel shows the mean ± SEM of average crawling speed (millimeters per second, mm/s) for day-1 adults from two independent experiments, n = ~55 worms for each genotype, *** p < 0.0001 (one-way ANOVA) compared to related control.

(C) pals-22 mutants have shorter lifespans compared to wild type or pals-22(PCR_R). Survival study was initiated with 96 worms (for each genotype) at L4 stage (day-0). Data shown is one represented trial of lifespan; two additional independent trials also show similar changes of lifespan in pals-22 mutants, p < 0.0001 (Log-rank test), comparing the wild type to mutants, or comparing pals-22(PCR_R) against pals-22(ot811).

(D) pals-22 mutants exhibit early accumulation of age pigment. The left panel shows representative pictures of age pigment (excitation: 364nm; emission: 380-420nm). The right panel shows the mean ± SEM of age pigment auto-fluorescence intensity of day-5 adults, n = 10 worms from each genotype, ns (not significant) or ** p < 0.01 (one-way ANOVA) compared to related control

Apart from the locomotory defects, we also noted abnormal survival of pals-22 mutants. Using standard lifespan assays, we observed premature death of pals-22 mutants, commending at about day 10 of adulthood (Fig. 8C). We also find that 5 day old adult animals display a signature change in aging, namely increased age pigment in the gut of adult worms (Fig. 8D). In light of these premature aging phenotypes, we surmise that the locomotory defects described above may also be an indication of premature aging.

DISCUSSION

Together with a parallel study by Reddy and colleagues (Reddyet al. 2017), our study provides the first functional characterization of the large, unusual family of pals genes in C. elegans. In the context of whole animal transcriptome profiling under different conditions, expression of members of the pals gene family has previously been shown to be induced upon various forms of cellular insults, ranging from exposure to intracellular fungal infections, to viral infection and to toxic compound exposure (Cuiet al. 2007; Bakowskiet al. 2014; Chenet al. 2017). We define here a function for one of the family members, pals-22 in controlling the silencing of repetitive DNA sequences. Even though the biochemical function of PALS proteins is presently unclear, the upregulation of many pals genes under conditions of cellular stress suggest that this gene family may be part of a host defense mechanism that protects animals/cells from specific insults. The C. elegans-specific expansion of pals genes may relate to their potential function in fending off species-specific stressors and/or encounters with species-specific pathogens. Consistent with the species-specificity of pals gene function, it has been noted that distinct wild isolates of C. elegans display an above-average accumulation of polymorphisms in most pals genes (Volkerset al. 2013).

The mutant phenotype of pals-22 as a transgene silencer, as well the dependence of this phenotype on small RNA production, indicates that PALS-22 may control gene expression via small RNA molecules. A role of PALS-22 in controlling gene expression is also illustrated in a parallel study in which pals-22 has been found to be required for the proper regulation of a battery of stress- and microspiridial infection-induced genes, including many of the pals genes themselves (Reddyet al. 2017). While the function of pals-22 in the RNAi process is not clear, there are numerous examples of mutants in which somatic transgene silencing is induced as a result of an increase in RNAi sensitivity, including rrf-3, eri-1, lin-35 and others (Simmeret al. 2002; Kennedyet al. 2004; Kimet al. 2005; Wanget al. 2005; Lehner et al. 2006; Fischeret al. 2008; Fischeret al. 2011; Fischeret al. 2013). Since the expression levels per copy of repetitive tandem arrays are much lower than for endogenous genes, transgene targeting siRNAs appear to be already abundant in wild-type transgenic strains, indicating a background level of transgene silencing in wild-type worms (Mello AND Fire 1995). These transgene-targeted siRNAs may be reduced in a mutant displaying enhanced transgene silencing (such as pals-22), perhaps because of a shift in the balance between the loading of transgene siRNAs into a silencing Argonaute (e.g. NRDE3) versus an anti-silencing Argonaute (e.g. CSR-1) (Shirayamaet al. 2012; Fischeret al. 2013).

The precise biochemical function of any PALS protein remains obscure. The only notable sequence relationship that we could find points to potential biochemical function of the PALS proteins in proteostasis. One of the LGIII clusters of pals genes (III: -21.90 cM) contains also a large number of fbxa genes, another vastly C. elegans-specific expanded gene family (Thomas 2006)(Fig̤ fbxa genes code for F-box proteins that are involved in protein degradation (Thomas 2006). One of the FBXA proteins in the LGIII cluster, FBXA-138, displays sequence similarities to a number of distinct PALS proteins within and outside the LGIII pals cluster, including PALS-22, PALS-23, PALS-32, PALS-25 and PALS-1. Even though PALS proteins are not predicted to contain a canonical F-box, it is conceivable that their distant sequence relationship to F box proteins may suggest a role of PALS proteins in protein degradation. How, in the case of PALS-22, such a function may relate to the control of gene expression via small RNA molecules is not clear.

Although it is difficult to unambiguously distinguish accelerated aging from general sickness, young adult pals-22 mutants clearly exhibit multiple features of aged animals—impaired mobility, elevated age pigments/lipofuscin, and shortened lifespan. Increased expression of repetitive sequences has been documented in aging C. elegans, Drosophila, and humans, and has been suggested to contribute to genomic instability and cell dysfunction (Sedivyet al. 2013); the physiological effect of decreases in the expression of repetitive sequences has not been explored. However, we note that other transgene silencing mutants (which are also RNAi hypersensitive), like rrf-3 and eri-1, do not display an aging defect (Zhanget al. 2009; Renet al. 2012). pals-22 may therefore be involved in novel aspects of small RNA-dependent gene silencing that may control the expression of genes involved in animal physiology.

ACKNOWLEDGEMENTS

We thank Chi Chen for the generation of transgenic strains, the CGC for providing strains (funded by NIH Office of Research Infrastructure Programs, P40 OD010440), Kevin Howe and Gary Williams at WormBase for their help obtaining and analyzing protein domain data, Sangeena Salam for initial life span assays, the Million Mutation Project for providing the pals-19(gk166606) and pals-25(gk891046) alleles, Emily Troemel for communicating unpublished results, Dylan Rahe for providing the GFP-tagged che-1(ot856) allele, HaoSheng Sun for providing the lin-4prom::yfp MiniMos strain, and members of the Hobert lab for comments on this manuscript. This work was supported by an R01AG046358 to MD, and a NJ Commission on Cancer Research Postdoctoral Fellowship DHFS16PPC070 to GW and the Howard Hughes Medical Institute (OH). E.L.-D. was funded by an EMBO long-term fellowship.

REFERENCES

  1. ↵
    Bakowski, M. A., C. A. Desjardins, M. G. Smelkinson, T. L. Dunbar, I. F. Lopez-Moyado et al. 2014 Ubiquitin-mediated response to microsporidia and virus infection in C. elegans. PLoS Pathog 10: e1004200.
    OpenUrlCrossRefPubMed
  2. ↵
    Brenner, S., 1974 The genetics of Caenorhabditis elegans. Genetics 77: 71–94.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    Celniker, S. E., L. A. Dillon, M. B. Gerstein, K. C. Gunsalus, S. Henikoff et al., 2009 Unlocking the secrets of the genome. Nature 459: 927–930.
    OpenUrlCrossRefPubMedWeb of Science
  4. ↵
    Celniker, S. E., D. A. Wheeler, B. Kronmiller, J. W. Carlson, A. Halpern et al., 2002 Finishing a whole-genome shotgun: release 3 of the Drosophila melanogaster euchromatic genome sequence. Genome Biol 3: RESEARCH0079.
  5. ↵
    Chen, K., C. J. Franz, H. Jiang, Y. Jiang and D. Wang, 2017 An evolutionarily conserved transcriptional response to viral infection in Caenorhabditis nematodes. BMC Genomics 18: 303.
    OpenUrlCrossRef
  6. ↵
    Chuong, E. B., N. C. Elde and C. Feschotte, 2017 Regulatory activities of transposable elements: from conflicts to benefits. Nat Rev Genet 18: 71–86.
    OpenUrlCrossRefPubMed
  7. ↵
    Cordaux, R., and M. A. Batzer, 2009 The impact of retrotransposons on human genome evolution. Nat Rev Genet 10: 691–703.
    OpenUrlCrossRefPubMedWeb of Science
  8. ↵
    Cui, M., and M. Han, 2007 Roles of chromatin factors in C. elegans development. WormBook: 1–16.
  9. ↵
    Cui, Y., S. J. McBride, W. A. Boyd, S. Alper and J. H. Freedman, 2007 Toxicogenomic analysis of Caenorhabditis elegans reveals novel genes and pathways involved in the resistance to cadmium toxicity. Genome Biol 8: R122.
    OpenUrlCrossRefPubMed
  10. ↵
    de Koning, A. P., W. Gu, T. A. Castoe, M. A. Batzer and D. D. Pollock, 2011 Repetitive elements may comprise over two-thirds of the human genome. PLoS Genet 7: e1002384.
    OpenUrlCrossRefPubMed
  11. Dickinson, D. J., A. M. Pani, J. K. Heppert, C. D. Higgins and B. Goldstein, 2015 Streamlined Genome Engineering with a Self-Excising Drug Selection Cassette. Genetics 200: 1035–1049.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    Doitsidou, M., N. Flames, A. C. Lee, A. Boyanov and O. Hobert, 2008 Automated screening for mutants affecting dopaminergic-neuron specification in C. elegans. Nat Methods 5: 869–872.
    OpenUrlCrossRefPubMedWeb of Science
  13. Doitsidou, M., N. Flames, I. Topalidou, N. Abe, T. Felton et al., 2013 A combinatorial regulatory signature controls terminal differentiation of the dopaminergic nervous system in C. elegans. Genes Dev 27: 1391–1405.
    OpenUrl
  14. ↵
    Doitsidou, M., R. J. Poole, S. Sarin, H. Bigelow and O. Hobert, 2010 C. elegans Mutant Identification with a One-Step Whole-Genome-Sequencing and SNP Mapping Strategy. PLoS ONE 5: e15435.
    OpenUrlCrossRefPubMed
  15. ↵
    Doolittle, W. F., and C. Sapienza, 1980 Selfish genes, the phenotype paradigm and genome evolution. Nature 284: 601–603.
    OpenUrlCrossRefPubMedWeb of Science
  16. ↵
    Edelstein, A., N. Amodaj, K. Hoover, R. Vale and N. Stuurman, 2010 Computer control of microscopes using microManager. Curr Protoc Mol Biol Chapter 14: Unit14 20.
    OpenUrl
  17. ↵
    Elbashir, S. M., W. Lendeckel and T. Tuschl, 2001 RNA interference is mediated by 21- and 22-nucleotide RNAs. Genes Dev 15: 188–200.
    OpenUrlAbstract/FREE Full Text
  18. ↵
    Finn, R. D., T. K. Attwood, P. C. Babbitt, A. Bateman, P. Bork et al., 2017 InterPro in 2017-beyond protein family and domain annotations. Nucleic acids research 45: D190–D199.
    OpenUrlCrossRefPubMed
    1. M. Chalfie and
    2. S. Kain.
    Fire, A., W. G. Kelly, M. Hsu, S. Xu, J. Ahnn et al., 1998 The Uses of Green Fluorescent Protein in Caenorhabditis elegans, pp. 153–168 in Green Fluorescent Protein. Properties, Applications and Protocols, edited by M. Chalfie and S. Kain. Wiley-Liss, New York.
  20. ↵
    Fischer, S. E., M. D. Butler, Q. Pan and G. Ruvkun, 2008 Trans-splicing in C. elegans generates the negative RNAi regulator ERI-6/7. Nature 455: 491–496.
    OpenUrlCrossRefPubMedWeb of Science
  21. ↵
    Fischer, S. E., T. A. Montgomery, C. Zhang, N. Fahlgren, P. C. Breen et al., 2011 The ERI-6/7 helicase acts at the first stage of an siRNA amplification pathway that targets recent gene duplications. PLoS Genet 7: e1002369.
    OpenUrlCrossRefPubMed
  22. ↵
    Fischer, S. E., Q. Pan, P. C. Breen, Y. Qi, Z. Shi et al., 2013 Multiple small RNA pathways regulate the silencing of repeated and foreign genes in C. elegans. Genes Dev 27: 2678–2695.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    Friedli, M., and D. Trono, 2015 The Developmental Control of Transposable Elements and the Evolution of Higher Species. Annual Review of Cell and Developmental Biology 31: 429–451.
    OpenUrlCrossRefPubMed
  24. Frokjaer-Jensen, C., M. W. Davis, M. Sarov, J. Taylor, S. Flibotte et al., 2014 Random and targeted transgene insertion in Caenorhabditis elegans using a modified Mos1 transposon. Nat Methods 11: 529–534.
    OpenUrlCrossRefPubMed
  25. ↵
    Gent, J. I., A. T. Lamm, D. M. Pavelec, J. M. Maniar, P. Parameswaran et al., 2010 Distinct phases of siRNA synthesis in an endogenous RNAi pathway in C. elegans soma. Mol Cell 37: 679–689.
    OpenUrlCrossRefPubMedWeb of Science
  26. ↵
    Gerstbrein, B., G. Stamatas, N. Kollias and M. Driscoll, 2005 In vivo spectrofluorimetry reveals endogenous biomarkers that report healthspan and dietary restriction in Caenorhabditis elegans. Aging Cell 4: 127–137.
    OpenUrlCrossRefPubMedWeb of Science
  27. ↵
    Goke, J., and H. H. Ng, 2016 CTRL+INSERT: retrotransposons and their contribution to regulation and innovation of the transcriptome. EMBO Rep 17: 1131–1144.
    OpenUrl
  28. ↵
    Goodier, J. L., and H. H. Kazazian, 2008 Retrotransposons revisited: the restraint and rehabilitation of parasites. Cell 135: 23–35.
    OpenUrlCrossRefPubMedWeb of Science
  29. ↵
    Grishok, A., J. L. Sinskey and P. A. Sharp, 2005 Transcriptional silencing of a transgene by RNAi in the soma of C. elegans. Genes Dev 19: 683–696.
    OpenUrlAbstract/FREE Full Text
  30. ↵
    Hacein-Bey-Abina, S., C. Von Kalle, M. Schmidt, M. P. McCormack, N. Wulffraat et al., 2003 LM02-associated clonal T cell proliferation in two patients after gene therapy for SCID-X1. Science 302: 415–419.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    Hadano, S., C. K. Hand, H. Osuga, Y. Yanagisawa, A. Otomo et al., 2001 A gene encoding a putative GTPase regulator is mutated in familial amyotrophic lateral sclerosis 2. Nature genetics 29: 166–173.
    OpenUrlCrossRefPubMedWeb of Science
  32. ↵
    Hobert, O., 2002 PCR fusion-based approach to create reporter gene constructs for expression analysis in transgenic C. elegans. Biotechniques 32: 728–730.
    OpenUrlCrossRefPubMedWeb of Science
  33. ↵
    Hsieh, J., J. Liu, S. A. Kostas, C. Chang, P. W. Sternberg et al., 1999 The RING finger/B-box factor TAM-1 and a retinoblastoma-like protein LIN-35 modulate context-dependent gene silencing in Caenorhabditis elegans. Genes Dev 13: 2958–2970.
    OpenUrlAbstract/FREE Full Text
  34. ↵
    Huelsenbeck, J. P., and F. Ronquist, 2001 MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics 17: 754–755.
    OpenUrlCrossRefPubMedWeb of Science
  35. ↵
    Jacobs, F. M. J., D. Greenberg, N. Nguyen, M. Haeussler, A. D. Ewing et al., 2014 An evolutionary arms race between KRAB zinc-finger genes ZNF91/93 and SVA/L1 retrotransposons. Nature 516: 242–245.
    OpenUrlCrossRefPubMedWeb of Science
  36. ↵
    Ji, N., and A. van Oudenaarden, 2012 Single molecule fluorescent in situ hybridization (smFISH) of C. elegans worms and embryos. WormBook: 1–16.
  37. ↵
    Kamath, R. S., and J. Ahringer, 2003 Genome-wide RNAi screening in Caenorhabditis elegans. Methods 30: 313–321.
    OpenUrlCrossRefPubMedWeb of Science
  38. ↵
    Kazazian, H. H., Jr.., 2004 Mobile elements: drivers of genome evolution. Science 303: 1626–1632.
    OpenUrlAbstract/FREE Full Text
  39. ↵
    Kelly, W. G., S. Xu, M. K. Montgomery and A. Fire, 1997 Distinct requirements for somatic and germline expression of a generally expressed Caernorhabditis elegans gene. Genetics 146: 227–238.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    Kennedy, S., D. Wang and G. Ruvkun, 2004 A conserved siRNA-degrading RNase negatively regulates RNA interference in C. elegans. Nature 427: 645–649.
    OpenUrlCrossRefPubMedWeb of Science
  41. Kerk, S. Y., P. Kratsios, M. Hart, R. Mourao and O. Hobert, 2017 Diversification of C. elegans Motor Neuron Identity via Selective Effector Gene Repression. Neuron 93: 80–98.
    OpenUrl
  42. ↵
    Kim, J. K., H. W. Gabel, R. S. Kamath, M. Tewari, A. Pasquinelli et al., 2005 Functional genomic analysis of RNA interference in C. elegans. Science 308: 1164–1167.
    OpenUrlAbstract/FREE Full Text
  43. ↵
    Lander, E. S., L. M. Linton, B. Birren, C. Nusbaum, M. C. Zody et al., 2001 Initial sequencing and analysis of the human genome. Nature 409: 860–921.
    OpenUrlCrossRefPubMedWeb of Science
  44. ↵
    Law, J. A., and S. E. Jacobsen, 2010 Establishing, maintaining and modifying DNA methylation patterns in plants and animals. Nat Rev Genet 11: 204–220.
    OpenUrlCrossRefPubMedWeb of Science
  45. ↵
    Le Thomas, A., A. K. Rogers, A. Webster, G. K. Marinov, S. E. Liao et al., 2013 Piwi induces piRNA-guided transcriptional silencing and establishment of a repressive chromatin state. Genes Dev 27: 390–399.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    Lehner, B., A. Calixto, C. Crombie, J. Tischler, A. Fortunato et al., 2006 Loss of LIN-35, the Caenorhabditis elegans ortholog of the tumor suppressor p105Rb, results in enhanced RNA interference. Genome Biology 7: R4.
    OpenUrlCrossRefPubMed
  47. ↵
    Lynch, M., and J. S. Conery, 2003 The origins of genome complexity. Science 302: 1401–1404.
    OpenUrlAbstract/FREE Full Text
  48. ↵
    Mello, C., and A. Fire, 1995 DNA transformation. Methods Cell Biol 48: 451–482.
    OpenUrlCrossRefPubMedWeb of Science
  49. ↵
    Mello, C. C., J. M. Kramer, D. Stinchcomb and V. Ambros, 1991 Efficient gene transfer in C.elegans: extrachromosomal maintenance and integration of transforming sequences. Embo J 10: 3959–3970.
    OpenUrlCrossRefPubMedWeb of Science
  50. ↵
    Mi, H., X. Huang, A. Muruganujan, H. Tang, C. Mills et al., 2017 PANTHER version 11: expanded annotation data from Gene Ontology and Reactome pathways, and data analysis tool enhancements. Nucleic Acids Res 45: D183–D189.
    OpenUrlCrossRefPubMed
  51. ↵
    Mi, H., A. Muruganujan, J. T. Casagrande and P. D. Thomas, 2013 Large-scale gene function analysis with the PANTHER classification system. Nat Protoc 8: 15511566.
    OpenUrl
  52. ↵
    Minevich, G., D. S. Park, D. Blankenberg, R. J. Poole and O. Hobert, 2012 CloudMap: A Cloud-Based Pipeline for Analysis of Mutant Genome Sequences. Genetics 192: 1249–1269.
    OpenUrlAbstract/FREE Full Text
  53. ↵
    Nowick, K., A. T. Hamilton, H. Zhang and L. Stubbs, 2010 Rapid Sequence and Expression Divergence Suggest Selection for Novel Function in Primate-Specific KRAB-ZNF Genes. Molecular Biology and Evolution 27: 2606–2617.
    OpenUrlCrossRefPubMedWeb of Science
  54. ↵
    Nussbaum-Krammer, C. I., M. F. Neto, R. M. Brielmann, J. S. Pedersen and R. I. Morimoto, 2015 Investigating the spreading and toxicity of prion-like proteins using the metazoan model organism C. elegans. J Vis Exp: 52321.
  55. ↵
    Orgel, L. E., and F. H. Crick, 1980 Selfish DNA: the ultimate parasite. Nature 284: 604–607.
    OpenUrlCrossRefPubMedWeb of Science
  56. ↵
    Parker, G. S., D. M. Eckert and B. L. Bass, 2006 RDE-4 preferentially binds long dsRNA and its dimerization is necessary for cleavage of dsRNA to siRNA. RNA (New York, N.Y.) 12: 807–818.
    OpenUrlAbstract/FREE Full Text
  57. ↵
    Quenneville, S., P. Turelli, K. Bojkowska, C. Raclot, S. Offner et al., 2012 The KRAB-ZFP/KAP1 System Contributes to the Early Embryonic Establishment of Site-Specific DNA Methylation Patterns Maintained during Development. Cell Reports 2: 766–773.
    OpenUrl
  58. ↵
    Reddy, K. C., T. Dror, J. Dowa, J. Panek, K. Chen et al., 2017 An intracellular pathogen response pathway promotes proteostasis in C. elegans BioRxiv submitted.
  59. ↵
    Ren, Y., S. Yang, G. Tan, W. Ye, D. Liu et al., 2012 Reduction of mitoferrin results in abnormal development and extended lifespan in Caenorhabditis elegans. PLoS One 7: e29666.
    OpenUrlCrossRefPubMed
  60. ↵
    Restif, C., C. Ibáñez-Ventoso, M. M. Vora, S. Guo, D. Metaxas et al., 2014 CeleST: computer vision software for quantitative analysis of C. elegans swim behavior reveals novel features of locomotion. PLoS computational biology 10: e1003702.
    OpenUrl
  61. ↵
    Ronquist, F., and J. P. Huelsenbeck, 2003 MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19: 1572–1574.
    OpenUrlCrossRefPubMedWeb of Science
  62. ↵
    Rowe, H. M., J. Jakobsson, D. Mesnard, J. Rougemont, S. Reynard et al., 2010 KAP1 controls endogenous retroviruses in embryonic stem cells. Nature 463: 237–240.
    OpenUrlCrossRefPubMedWeb of Science
  63. ↵
    Schlesinger, S., and S. P. Goff, 2015 Retroviral transcriptional regulation and embryonic stem cells: war and peace. Molecular and cellular biology 35: 770–777.
    OpenUrlAbstract/FREE Full Text
  64. ↵
    Schneider, C. A., W. S. Rasband and K. W. Eliceiri, 2012 NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9: 671–675.
    OpenUrlCrossRefPubMedWeb of Science
  65. ↵
    Sedivy, J. M., J. A. Kreiling, N. Neretti, M. De Cecco, S. W. Criscione et al., 2013 Death by transposition - the enemy within? Bioessays 35: 1035–1043.
    OpenUrlCrossRefPubMed
  66. Serrano-Saiz, E., R. J. Poole, T. Felton, F. Zhang, E. D. de la Cruz et al., 2013 Modular Control of Glutamatergic Neuronal Identity in C. elegans by Distinct Homeodomain Proteins. Cell 155: 659–673.
    OpenUrlCrossRefPubMedWeb of Science
  67. ↵
    Shapiro, J. A., and R. von Sternberg, 2005 Why repetitive DNA is essential to genome function. Biological reviews of the Cambridge Philosophical Society 80: 227–250.
    OpenUrlCrossRefPubMed
  68. ↵
    Shirayama, M., M. Seth, H.-C. Lee, W. Gu, T. Ishidate et al., 2012 piRNAs initiate an epigenetic memory of nonself RNA in the C. elegans germline. Cell 150: 65–77.
    OpenUrlCrossRefPubMedWeb of Science
  69. ↵
    Simmer, F., M. Tijsterman, S. Parrish, S. Koushika, M. Nonet et al., 2002 Loss of the Putative RNA-Directed RNA Polymerase RRF-3 Makes C. elegans Hypersensitive to RNAi. Curr Biol 12: 1317.
    OpenUrlCrossRefPubMedWeb of Science
  70. ↵
    Stefanakis, N., I. Carrera and O. Hobert, 2015 Regulatory Logic of Pan-Neuronal Gene Expression in C. elegans. Neuron 87: 733–750.
    OpenUrlCrossRefPubMed
  71. ↵
    Stein, L. D., Z. Bao, D. Blasiar, T. Blumenthal, M. R. Brent et al., 2003 The genome sequence of Caenorhabditis briggsae: a platform for comparative genomics. PLoS Biology 1: E45.
    OpenUrlCrossRefPubMed
  72. ↵
    Tabara, H., M. Sarkissian, W. G. Kelly, J. Fleenor, A. Grishok et al., 1999 The rde-1 gene, RNA interference, and transposon silencing in C. elegans. Cell 99: 123–132.
    OpenUrlCrossRefPubMedWeb of Science
  73. ↵
    Tabara, H., E. Yigit, H. Siomi and C. C. Mello, 2002 The dsRNA binding protein RDE-4 interacts with RDE-1, DCR-1, and a DExH-box helicase to direct RNAi in C. elegans. Cell 109: 861–871.
    OpenUrlCrossRefPubMedWeb of Science
  74. ↵
    Thomas, J. H., 2006 Adaptive evolution in two large families of ubiquitin-ligase adapters in nematodes and plants. Genome Res 16: 1017–1030.
    OpenUrlAbstract/FREE Full Text
  75. ↵
    Thompson, O., M. Edgley, P. Strasbourger, S. Flibotte, B. Ewing et al., 2013 The million mutation project: a new approach to genetics in Caenorhabditis elegans. Genome Res 23: 1749–1762.
    OpenUrlAbstract/FREE Full Text
  76. ↵
    Toth, K. F., D. Pezic, E. Stuwe and A. Webster, 2016 The piRNA Pathway Guards the Germline Genome Against Transposable Elements. Adv Exp Med Biol 886: 51–77.
    OpenUrlCrossRef
  77. ↵
    Tseng, R. J., K. R. Armstrong, X. Wang and H. M. Chamberlin, 2007 The bromodomain protein LEX-1 acts with TAM-1 to modulate gene expression in C. elegans. Mol Genet Genomics 278: 507–518.
    OpenUrlCrossRefPubMed
  78. ↵
    Vasale, J. J., W. Gu, C. Thivierge, P. J. Batista, J. M. Claycomb et al., 2010 Sequential rounds of RNA-dependent RNA transcription drive endogenous small-RNA biogenesis in the ERGO-1/Argonaute pathway. Proc Natl Acad Sci U S A 107: 3582–3587.
    OpenUrlAbstract/FREE Full Text
  79. ↵
    Volkers, R. J., L. B. Snoek, C. J. Hubar, R. Coopman, W. Chen et al., 2013 Gene-environment and protein-degradation signatures characterize genomic and phenotypic diversity in wild Caenorhabditis elegans populations. BMC Biol 11: 93.
    OpenUrlCrossRefPubMed
  80. ↵
    Wallace, I. M., O. O’Sullivan, D. G. Higgins and C. Notredame, 2006 M-Coffee: combining multiple sequence alignment methods with T-Coffee. Nucleic Acids Res 34: 1692–1699.
    OpenUrlCrossRefPubMedWeb of Science
  81. ↵
    Wang, D., S. Kennedy, D. Conte, Jr.., J. K. Kim, H. W. Gabel et al., 2005 Somatic misexpression of germline P granules and enhanced RNA interference in retinoblastoma pathway mutants. Nature 436: 593–597.
    OpenUrlCrossRefPubMedWeb of Science
  82. ↵
    Wolf, D., and S. P. Goff, 2009 Embryonic stem cells use ZFP809 to silence retroviral DNAs. Nature 458: 1201–1204.
    OpenUrlCrossRefPubMedWeb of Science
  83. ↵
    Zamore, P. D., T. Tuschl, P. A. Sharp and D. P. Bartel, 2000 RNAi: double-stranded RNA directs the ATP-dependent cleavage of mRNA at 21 to 23 nucleotide intervals. Cell 101: 25–33.
    OpenUrlCrossRefPubMedWeb of Science
  84. Zhang, L., J. D. Ward, Z. Cheng and A. F. Dernburg, 2015 The auxin-inducible degradation (AID) system enables versatile conditional protein depletion in C. elegans. Development 142: 4374–4384.
    OpenUrlAbstract/FREE Full Text
  85. ↵
    Zhang, M., M. Poplawski, K. Yen, H. Cheng, E. Bloss et al., 2009 Role of CBP and SATB-1 in aging, dietary restriction, and insulin-like signaling. PLoS Biol 7: e1000245.
    OpenUrlCrossRefPubMed
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Posted June 02, 2017.
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pals-22, a member of an expanded C. elegans gene family, controls silencing of repetitive DNA
Eduardo Leyva-Díaz, Nikolaos Stefanakis, Inés Carrera, Lori Glenwinkel, Guoqiang Wang, Monica Driscoll, Oliver Hobert
bioRxiv 145094; doi: https://doi.org/10.1101/145094
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