Transposon Regulation in the Caenorhabditis elegans Germline and Soma

Transposon Regulation in the Caenorhabditis elegans Germline and Soma | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Transposon Regulation in the Caenorhabditis elegans Germline and Soma View ORCID Profile Cindy Chang , Dong Cao , Daniel J. Pagano , View ORCID Profile Scott Kennedy doi: https://doi.org/10.1101/2025.10.09.681459 Cindy Chang 1 Department of Genetics, Harvard Medical School , Boston, MA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Cindy Chang Dong Cao 1 Department of Genetics, Harvard Medical School , Boston, MA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Daniel J. Pagano 1 Department of Genetics, Harvard Medical School , Boston, MA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Scott Kennedy 1 Department of Genetics, Harvard Medical School , Boston, MA, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Scott Kennedy For correspondence: kennedy{at}genetics.med.harvard.edu Abstract Full Text Info/History Metrics Supplementary material Data/Code Preview PDF Abstract Transposons are parasitic nucleic acids present in most genomes. The ability of transposons to mobilize makes them a source of genetic diversity and a threat to genome integrity. Interestingly, mutations in the C. elegans gene mut-2/rde-3 increases the rate of transposition in the germline, but not in the soma, suggesting that the C. elegans germline and soma employ different strategies to regulate Tc1 transposition. Here, we develop fluorescence reporters for studying DNA transposon regulation in living C. elegans in a tissue-specific manner and we use candidate gene testing and genetic screening approaches to identify factors that regulate Tc1 mobility in the germline and/or the soma of the animal. We find that both cytoplasmic and nuclear components of the RNA interference (RNAi) pathway silence Tc1 in the germline, but not in the soma. We identify a novel pathway involving the C. elegans ortholog of HNRNPC, and a gene we term suppressor of transposon mobilization ( stm ) -1 , which regulates Tc1 primarily in the soma, likely by binding Tc1 RNA and preventing its splicing. Our findings reveal tissue-specific strategies for regulating parasitic nucleic acids and pave the way for future studies exploring how and why different tissues adopt different transposon silencing systems. Introduction Transposons are mobile genetic elements present in almost all eukaryotic genomes, where they often represent a large fraction of the genome (3-85%) [ 1 ]. For example, 40-50% of the human genome is composed of active transposons or inactive transposon relics [ 1 ]. Transposon insertions can disrupt host-gene functions, and high-copy transposons make genomes vulnerable to chromosome rearrangements [ 2 ]. To limit these deleterious effects of transposons, organisms have evolved defense mechanisms, including DNA, chromatin, and small RNA-based systems, which collectively silence transposon expression and restrict transposon mobility [ 3 – 6 ]. Transposons can be divided into two classes; The retrotransposons and the DNA transposons. Retrotransposons use an RNA-intermediate, which is subjected to reverse transcription, prior to transposition. Retrotransposons replicate via an RNA intermediate and are the most abundant type of transposon in the human genome, where they comprise >40% of the genome [ 7 ]. DNA transposons employ a copy-and-paste mechanism for transposition. DNA transposons are inactive in the human genome, yet relics of ancient DNA transposons comprise (∼2-3%) of the genome [ 7 ]. In the C. elegans genome, DNA transposons are the more common transposon, comprising 12% of the genome [ 8 , 9 ]. The most active and abundant of these elements are the Tc1/mariner class DNA transposons [ 9 ]. Tc1/mariner DNA transposons typically produce a single RNA that, after splicing and polyadenylation, is translated into transposase, which encodes the enzyme responsible for transposon excision and re-insertion [ 10 , 11 ]. The evolutionary impacts of transposition in the germline and soma differ for transposons and their hosts. For transposons, germline transposition is necessary for survival, while somatic transposition is likely of little to no benefit as it does not lead to a heritable change in copy number [ 12 ]. For hosts, the short-term fitness costs associated with transposition in the germline or soma are typically negative because transposition is more likely to disrupt, rather than improve, cell functions. Indeed, transposition in both the germline and soma have been linked to disease in humans [ 13 – 16 ]. And the impacts of transposition in the germline are likely to be more severe for the host than transposition in the soma because germline transposition impacts all cells (germ and soma) and these impacts are heritable. The differing goals of transposons and their host cells, as well as the differing impacts of transposition in the soma or the germline, suggest that the strategies employed by multicellular organisms to regulate their transposons may differ in the germline and soma. Consistent with this idea, in the Drosophila soma and germline, DNA transposons termed P elements are regulated by distinct alternative splicing programs [ 17 ]. Additionally, the Gypsy retrotransposon is inhibited by the piRNA flamenco locus in somatic cells of the germline but not in germ cells themselves [ 18 ]. In C. elegans , the rate at which the Mariner class DNA transposon Tc1 transposes is thought to be 1000x higher in the soma than in the germline, and loss of the RNA interference (RNAi) factor RDE-3/MUT-2 increases the rate of Tc1 transposition in the germline but not in the soma [ 19 – 21 ]. These observations hint that, as in flies, distinct mechanisms may exist to regulate transposons in the germline and soma of C. elegans [ 21 ]. Heterogenous nuclear ribonucleoprotein C (HNRNPC) is an abundant, nuclear protein that associates with most pre-mRNA [ 22 , 23 ]. HNRNPC regulates many aspects of gene expression, including pre-mRNA splicing as well as cleavage and polyadenylation [ 23 – 25 ]. Transposons often harbor cryptic splice sites, which can lead to aberrant splicing of transposon sequences into host mRNAs, a process referred to as transposon exonization. In mammals, HNRNPC prevents the exonization of Alu retrotransposons, which are the most abundant retrotransposon in the human genome [ 26 , 27 ]. Current models posit that HNRNPC binds uridine-rich tracts in Alu RNA [ 23 , 27 – 33 ] to prevent the splicing factor U2AF65 from binding these elements, thus inhibiting Alu exonization [ 27 ]. Whether the role of HNRNPC in transposon regulation is conserved or whether the pathway is differentially employed in soma/germline contexts is not yet known. Here we report the development of reporter genes for visualizing Tc1 excision in the soma and germline of living C. elegans . We use these reporters to identify cytoplasmic and nuclear RNA interference (RNAi) factors that limit Tc1 excision in the germline, but not in the soma. Additionally, we use a genetic screening approach to identify two nuclear-localized proteins, including the C. elegans ortholog of HNRNPC, HRPC-1, which we show are important for repressing Tc1 in the soma, likely by binding and preventing the splicing of Tc1 RNA. Thus, the C. elegans germline and soma employ distinct strategies to control their transposons; RNAi in the germline; and an HNRNPC-based pathway in the soma. Results Reporter genes to visualize transposon excision in C. elegans C. elegans Tc1 is a Mariner class DNA transposon which is mobile in the C. elegans soma and germline [ 10 ]. To explore the biology of transposons in an intact animal system, we engineered a reporter gene under the control of an ubiquitous promoter ( eft-3 p) to drive expression of mScarlet-I in all cells of the C. elegans soma and germline. A Tc1 element was then inserted 6 nucleotides upstream of the eft-3 p ::mScarlet-I start codon ( Figure 1A ) with the expectation that: 1) the transposon would likely interfere with mScarlet-I expression; 2) transposon excision would permit mScarlet-I expression in those cells in which Tc1 had excised; and 3) this would allow us to monitor transposon activity in all cells and during all stages of development of a living animal. We used the CRISPR/Cas9 system to insert the eft-3 p:: Tc1::mScarlet-I reporter into the C. elegans genome. A control mScarlet-I reporter gene lacking a Tc1 element was inserted into the same chromosomal site. Fluorescence imaging revealed that, as expected; 1) mScarlet-I was expressed in most/all cells in all eft-3 p ::mScarlet-I animals; and 2) the addition of the Tc1 element inhibited this mScarlet-I expression ( Figure 1B ). In 77% of eft-3 p:: Tc1::mScarlet-I animals, no mScarlet-I was observed in any cell of the germline or soma ( Figure 1B ). In 23% of animals, 1-2 somatic cells expressed mScarlet-I ( Figure 1B and see below). Download figure Open in new tab Figure 1. Reporter genes designed to detect Tc1 excision in living animals. (A) Schematic of Tc1::mScarlet-I reporter gene engineered to monitor Tc1 excision. NLS, nuclear localization signal; UTR, untranslated region. Rightward arrow indicates transcription start site and direction of transcription. (B) (Top) Fluorescence micrograph of representative eft-3 p ::mScarlet-I ( mScarlet-I ) animal. Percentage indicates percentage of eft-3 p ::mScarlet-I animals expressing mScarlet-I in all cells. (Middle, Bottom) Fluorescence micrographs of representative eft-3 p :: Tc1::mScarlet-I ( Tc1::mScarlet-I ) animals. 77% of Tc1::mScarlet-I animals do not show any expression of mScarlet-I; 23% of Tc1::mScarlet-I animals have somatic expression of mScarlet-I. Magnification of mScarlet-I positive cell (arrow) is shown. (C) Germline excision of Tc1 from Tc1::mScarlet-I in rde-3(-) animal. mScarlet-I expression is observed in all cells of the germline and soma. Percentage indicates the rate of germline excision from Tc1::mScarlet-I in rde-3(-) animals. During this study, no germline Tc1 excision was ever observed in wild-type animals. (D) Sanger sequencing of Tc1 excision events from PCR on DNA isolated from glp-1 ; Tc1::mScarlet-I animals raised at 25℃, which ablates most germ cells (see Materials and Methods). The PCR strategy allows relatively rare Tc1 excision events in the soma to be amplified because full-length Tc1 containing DNA is poorly amplified by the PCR reaction. (E) (Top) Schematic of Tc1::sfgfp reporter gene engineered to monitor Tc1 excision. (Middle) Somatic excision from Tc1::sfgfp in wild-type animals (arrow, somatic cell expressing sfGFP). (Bottom) Germline excision of Tc1 from Tc1::sfgfp in rde-3(-) background. Tc1 reporter genes recapitulate transposon behaviors We asked if the Tc1::mScarlet-I reporter gene recapitulated known properties of Tc1 and, therefore, could be used to study transposon biology in a living animal. Tc1 excision in somatic cells would be expected to produce animals expressing fluorescent protein in one or more somatic cells. As outlined above, we observed that 23% of Tc1::mScarlet-I animals expressed high levels of mScarlet-I in one or more somatic cells, which suggests that Tc1 had excised in these cells ( Table 1 and Figure 1B ). The rate of putative Tc1 excision in the soma was determined by dividing the total number of animals possessing one or more mScarlet-I positive cells (1304) by the total number of somatic cells scored (≅4.91×10 5 ) ( Table 1 ). This analysis set the somatic excision rate of Tc1::mScarlet-I at ≥ 2.7 x10 -4 , which is consistent with previous reports for endogenous Tc1 element activity in the soma (approximately 10 -3 ) [ 19 , 34 ]. Four additional lines of evidence show that the Tc1::mScarlet-I reporter gene recapitulates known aspects of Tc1 biology in the C. elegans soma and germline. View this table: View inline View popup Download powerpoint Table 1. Somatic Tc1 excision rates. Somatic Tc1 excision events were quantified by counting the number of somatic cells expressing mScarlet-I (indicated as mScarlet-I (+)) using the Tc1::mScarlet-I reporter gene. Somatic Tc1 excision rates were estimated using the following formula: (# of animals with mScarlet-I(+) cells) x (# of mScarlet-I(+) cells per animal) / ((total # of animals scored) x (1000 = # somatic cells/animal)). For wild-type, if an animal had any mScarlet-I (+) cells the animal typically (73%) had only one mScarlet-I(+) cell. The remaining animals had two (10%) or three (13%) mScarlet-I(+) cells. For these animals, the number of mScarlet-I(+) cells per animal was set as 1 in the above calculation, and the “≥” symbol was used to account for animals that had more than one mScarlet-I(+) cell. For hrpc-1 and stm-1 mutants, all animals showed Tc1 excision and the number of mScarlet-I(+) cells per animal was 37.4 and 26.5 respectively (See Fig 3B ). Data are aggregates from at least 3 independent experiments per genotype. Tc1::mSc , Tc1::mScarlet-I . First, Tc1 excision is reported to be ∼1000 fold lower in the C. elegans germline than in the soma [ 19 , 35 , 36 ]. The rate of germline excision for Tc1 in wild-type C. elegans germ cells is reportedly ∼10 -7 [ 37 ]. Tc1 excision from Tc1::mScarlet-I in germ cells would be expected to result in animals that express fluorescent protein in all cells of the germline and soma and this expression state should be heritable. Tc1::mScarlet-I germline excision rates were determined by growing animals to high density over several generations and then determining the percentage of plates containing one or more animals expressing mScarlet-I in all cells of the soma and germline, which is indicative of germline transposition. A Poisson distribution method (see Materials and Methods) was used to determine maximum excision rates. Using this approach, we calculated a germline excision rate for Tc1::mScarlet-I animals of <2.2×10 -7 ( Table 2 ), which is similar to the published rate of ∼10 -7 for endogenous Tc1 elements [ 37 ]. View this table: View inline View popup Download powerpoint Table 2. Germline Tc1 excision rates. Germline Tc1 excision events were detected by counting animals in Tc1::mScarlet-I or Tc1::sfgfp backgrounds (specified in “Reporter used” column) that heritably express mScarlet-I or sfGFP in all cells. Animals were singled onto plates and allowed to grow until plates were populated by F1 or F2/3 broods as indicated. Populations were scored for presence or absence of one or more animals expressing fluorescence in all cells, indicating at least one germline excision event. For genotypes with high germline excision rates ( rde-3(ne3370) , mut-16(pk710) , deps-1(bn124) ), plates were scored in the F1 generation. For genotypes with low or undetectable germline excision rates (wild-type, mut-16(mg461) , znfx-1(ne4583) , nrde-2(gg873), hrpc-1(gg909), stm-1(gg893) ), plates were scored in the F2/3 generations. Tc1::sfgfp animals were all scored in F2/3 generations as the Tc1 excision excision rate from this reporter is lower than from Tc1::mScarlet-I. The proportion of plates for each genotype showing germline excision event(s) and the average number of animals per plate at the time of scoring for each genotype were used to calculate the germline excision rate using a Poisson distribution method (see Materials and Methods for formula explanation). For genetic backgrounds where the rate of germline excision was so low that no excision events were ever observed, the Poisson distribution method was used to set a maximum rate of germline excision (indicated with “<”). Data are aggregates from at least 3 independent experiments per genotype. ND, not determined. Tc1::mSc , Tc1::mScarlet-I . gg1160 and gg1161 are independent alleles created by CRISPR/Cas9, both equivalent to the ne298 allele [ 42 , 61 ]. Second, rde-3 and mut-16 limit Tc1 activity in the germline of C. elegans [ 36 , 38 ]. rde-3(-); Tc1::mScarlet-I and mut-16(pk710); Tc1::mScarlet-I animals exhibited a >1000 fold increase in the rate of Tc1 excision from Tc1::mScarlet-I in the germline as indicated by the appearance of animals expressing mScarlet-I in all cells ( Table 2 and Figure 1C ). Thus, excision of Tc1 from Tc1::mScarlet-I is regulated by factors that regulate endogenous Tc1 elements in the germline. Additionally, RDE-3 is reported to suppress excision of Tc1 in the germline, but not in the soma [ 36 ]. We observed that RDE-3(-) animals exhibited increased Tc1 excision from Tc1::mScarlet-I in the germline (see above) and did not show increase rates of Tc1 excision from Tc1::mScarlet-I in the soma ( Table 1 ). Third, 78% of the time endogenous Tc1 elements, which insert into TA dinucleotides, leave behind a four base pair TATGTA or TACATA excision footprint when excised from somatic cells [ 19 ]. We cloned and sequenced eleven independent somatic Tc1::mScarlet-I excision events observed in Tc1::mScarlet-I animals and in 8/11 cases (73%), we observed a TATGTA or TACATA four base pair insertion at the TA dinucleotide formerly occupied by Tc1 in Tc1::mScarlet-I ( Figure 1D ). Finally, we constructed a second Tc1 reporter gene in which a Tc1 element was inserted into the coding sequence of a superfolder green fluorescent protein ( sfgfp) gene driven under the control of a distinct, ubiquitous promoter ( rpl-28 p) ( Figure 1E ). The reading frame of Tc1::sfgfp was engineered such that a typical somatic Tc1 excision footprint (+4) would enable in-frame sfGFP translation ( Figure 1E ). Tc1::sfgfp behaved like Tc1::mScarlet-I , exhibiting properties previously ascribed to endogenous Tc1 elements ( Figure 1E ). These properties included elevated germline Tc1 excision rates in rde-3(-) animals ( Table 2 ). For unknown reasons, the germline excision rate from Tc1::sfgfp in rde-3(-) animals occurred at a ≅10x lower rate than that of Tc1::mScarlet-I ( Table 2 ). Together, these data establish that Tc1 reporter genes recapitulate known properties of endogenous C. elegans Tc1 elements and, therefore, can be used to explore transposon biology in an intact animal system. Identification of factors silencing Tc1 in the germline C. elegans RDE-3 and MUT-16 prevent Tc1 mobilization in the germline [ 21 , 38 ]. rde-3 encodes a poly(UG) nucleotidyltransferase [ 39 , 40 ] and mut-16 encodes a low complexity Q/N-rich protein [ 38 ]. Both RDE-3 and MUT-16 localize within germ cells to non-membrane enclosed cytoplasmic organelles termed Mutator foci ( Figure 2A ), where they contribute to the biogenesis of endogenous (endo) small interfering (si)RNAs, which are thought to negatively regulate the Tc1 transposase mRNA, thereby preventing Tc1 transposition [ 41 – 44 ]. Using our Tc1::mScarlet-I reporter gene, we observed, as expected, that mut-16 suppresses germline Tc1 excision from Tc1::mScarlet-I in the germline ( Table 2 and Figure 2B ). Next, we used the Tc1::mScarlet-I reporter to ask if other siRNA-related genes and pathways might coordinate with RDE-3 and MUT-16 to silence transposons in the C. elegans germline. Mutator foci localize near the outer membrane of germ cell nuclei juxtaposed to nuclear pores and directly adjacent to two other perinuclear subcompartments of the germ granule termed the P granule and the Z granule [ 44 , 45 ] ( Figure 2A ). Animals lacking DEPS-1, which is a low-complexity protein that localizes to P granules and which contributes to P granule formation [ 46 ], exhibited an increase in germline Tc1 excision from Tc1::mScarlet-I ( Figure 2B , Table 2 ). Animals lacking the helicase ZNFX-1, which localizes to Z granules-where it binds mRNAs undergoing siRNA-based gene silencing [ 47 ] - exhibited a moderate increase in germline Tc1 excision, which became evident after lineages were grown to high densities for two to three generations ( Figure 2C , Table 2 ). The data suggests that the various perinuclear germ granule compartments help coordinate anti-transposon defenses in C. elegans , perhaps by concentrating siRNA pathway proteins near nuclear pores to surveille transposon mRNAs exiting nuclei [ 48 ]. In addition to these cytoplasmic functions, C. elegans siRNAs also regulate gene expression within nuclei by modifying chromatin states and inhibiting transcription elongation (termed nuclear RNAi) ( Figure 2A ) [ 49 , 50 ]. We asked if the nuclear RNAi pathway might contribute to transposon silencing in the germline. Indeed, animals lacking NRDE-2, which is an RNA binding protein required for nuclear RNAi [ 51 ] exhibited a moderate increase in germline Tc1 excision, which was evident after two to three generations of growth ( Figure 2C , Table 2 ). We conclude that the cytoplasmic and nuclear branches of the RNAi system coordinately limit transposon excision in the C. elegans germline. Download figure Open in new tab Figure 2. Germline and soma employ different mechanisms to regulate Tc1 excision. (A) Schematic of the C. elegans RNAi pathway showing factors assayed for roles in suppressing Tc1 excision, including the germ granule components DEPS-1 (located in the P granule), ZNFX-1 (in the Z granule), RDE-3 and MUT-16 (in Mutator foci), and the nuclear RNAi factor NRDE-2, which inhibits gene expression at the level of transcription. ( B and C ) Percentage of plates containing one or more animals with germline Tc1 excision (indicated by mScarlet-I expression in all cells) when populations started with a single animal were expanded to the F1 generation (B) or F2/F3 generations (C) . Total plates scored per genotype per replicate were 6-19. ≥3 replicates scored per genotype. Error bars denote mean ± standard deviation (SD). WT, wild-type. (D) Somatic Tc1 excision rates visualized as percentage of progeny animals expressing mScarlet-I in one or more somatic cells from independently singled animals. Data points represent independent lineages and are from ≥3 replicates. Total lineages scored per genotype were 25-116. WT is wild-type. ns, not significant; **, p≤ 0.01; ***, p ≤ 0.001 (Kruskal-Wallis test with Dunn’s test for multiple comparisons). Error bars denote mean ± SD. Germline anti-transposon systems do not regulate Tc1 in the soma RDE-3, MUT-16, and NRDE-2, which silence transposons in the germline (see above), are expressed and/or are active in both the germline and the soma [ 40 , 43 , 51 ]. Therefore, we asked if these inhibitors of germline Tc1 might also inhibit Tc1 excision in the soma. Surprisingly, presumed null mutations in rde-3 , deps-1 , znfx-1 , or nrde-2 did not increase Tc1 excision in the soma ( Figure 2D , Table 1 ) as they did in the germline. For unknown reasons, rde-3 , mut-16(pk710) , and nrde-2 mutants actually exhibited a subtle, yet statistically significant, decrease in somatic excision of Tc1 from the Tc1::mScarlet-I reporter gene ( Figure 2D ). The mut-16(pk710) allele ablates MUT-16 function in both the soma and germline while the mut-16(mg461) allele deletes a portion of the mut-16 promoter, which abolishes MUT-16 function in the soma, but not the germline [ 43 ]. While mut-16(pk710) animals showed elevated rates of Tc1 excision in the germline, somatic excision rates were not elevated ( Figure 2D , Table 1 ). The soma-specific allele mut-16(mg461) failed to alter excision rates in either the germline or the soma ( Figures 2C-D , Tables 1 - 2 ). Together, the data show that the germline RNAi-based anti-transposon system does not limit Tc1 excision in the C. elegans soma, despite components of these pathways being present and active in this tissue. Identification of genes that limit Tc1 excision in the soma We wondered if other, currently unknown systems might regulate Tc1 in the soma. To identify such systems, we conducted a forward genetic screen using Tc1::mScarlet-I; Tc1::sfgfp double reporter animals to identify mutations that increased the frequency of Tc1 excision from both reporters in somatic cells (Figure S1A). The screen identified 21 mutant strains exhibiting >3x more somatic cells expressing mScarlet-I and sfGFP than wild-type animals. Seven of these 21 mutant strains exhibited a dramatic >60x elevation in somatic Tc1 excision as well as a delay in developmental timing (see below). We focused our efforts on characterizing these seven alleles due to the high expressivity of their elevated Tc1 excision phenotypes and because their shared developmental pleiotropy hinted at shared molecular function. HRPC-1 and STM-1 limit Tc1 excision in the soma Positional mapping and whole genome sequencing identified candidate mutations in all seven strains. In six of the seven strains, we identified mutations in the gene z inc finger putative transcription factor ( ztf)-4 ( Figure 3A ), which were each associated with a ≅85x elevation in Tc1 somatic excision (mean number of mScarlet-I positive cells per animal: 0.44 in wild-type and 37.4 in mutant) ( Figure 3B ). C. elegans ztf-4 is predicted to encode one zinc finger and an RNA recognition motif (RRM) domain ( Figure 3A ). Basic Local Alignment Search Tool (BLAST) analyses suggested that ZTF-4 is orthologous to vertebrate heterogeneous nuclear ribonucleoprotein C (HNRNPC) protein, with homology most pronounced in the RRM domain (Figure S1B). For clarity, we renamed ztf-4 as h ete r ogeneous nuclear p rotein C -like 1 ( hrpc-1) . All six hrpc-1 mutations identified by the genetic screen encoded missense alleles, four of which altered amino acids within the RRM domain of HRPC-1 ( Figures 3A and S1B). CRISPR/Cas9-based mutagenesis introducing the A132V RRM mutation into wild-type hrpc-1 resulted in animals that exhibited elevated levels of somatic Tc1 excision from Tc1::mScarlet-I and Tc1::sfgfp , confirming that hrpc-1 mutation causes an enhanced somatic Tc1 excision phenotype ( Figure 3C ). CRISPR/Cas9-based deletion of >90% of the hrpc-1 locus also resulted in animals exhibiting high levels of somatic Tc1 excision (Figure S2), indicating that one function of HRPC-1 in C. elegans is to limit Tc1 excision in somatic cells, and suggesting that the six hrpc-1 alleles identified in our genetic screen were loss-of-function. Download figure Open in new tab Figure 3. HRPC-1 and STM-1 regulate Tc1 primarily in the soma. (A) ztf-4 / hrpc-1 and F54D8.6 / stm-1 alleles identified in the genetic screen (“EMS allele(s)”) or generated by CRISPR-Cas9 (“CRISPR allele”) are shown. Unless otherwise indicated, hrpc-1 and stm-1 alleles used in this work are the CRISPR-Cas9-derived mutations that lead to HRPC-1(A132V) or STM-1(g.19_22>TAACTAGT). a.a., amino acid; RRM, RNA recognition motif; ZF, zinc finger domain. (B) Quantification of somatic excision events ((mScarlet-I (+) somatic cells)) in Tc1::mScarlet-I expressing strains of the indicated genotypes. n=51-59 animals per genotype. Data are from 3 biological replicates. ***, p ≤ 0.001 (Kruskal-Wallis test with Dunn’s test for multiple comparisons). Error bars denote mean ± SD. WT, wild-type. (C) Fluorescent micrographs of the anterior (head) of Tc1::mScarlet-I or Tc1::sfgfp animals of indicated genotypes showing that the number of sfGFP or mScarlet (+) cells is elevated in strains of the indicated genotypes. Scale bar, 30 μm. (D) (Top) Schematic of qPCR-based assay to quantify endogenous Tc1 excision events from genomic DNA. qPCR settings were optimized such that amplicons are detected only if Tc1 is excised. (Bottom) Quantification of Tc1 excision in hrpc-1 or stm-1 for two endogenous Tc1 sites ( Tc1#20 : WBTransposon00000020; Tc1#33 : WBTransposon00000033). Fold changes in excision rate were normalized to mean value in wild-type. n=12 per genotype, from 3 biological replicates. ***, p ≤ 0.001 (one-way ANOVA with Dunnett’s multiple comparisons test). Mean ± SD. WT, wild-type. (E) qPCR quantification of Tc1 excision for Tc1#20 in strains with or without auxin-induced depletion of HRPC-1 in the germline or soma (see Methods). n=11-12 per genotype/treatment condition, from 3 biological replicates. ns, not significant; ***, p ≤ 0.001 (Student’s t test). Mean ± SD. In the remaining mutant strain, which did not harbor a hrpc-1 mutation, and which exhibited an ≅60x elevation in Tc1 somatic excision (mean number of mScarlet-I positive cells per animal: wild-type, 0.44; mutant, 26.5) ( Figure 3B ), genome sequencing and positional mapping identified a candidate mutation in the gene F54D8.6 ( Figure 3A ). CRISPR/Cas9-based introduction of a stop codon, or a short insertion that creates a frameshift at the 7th amino acid of F54D8.6 , resulted in animals exhibiting high rates of Tc1 excision from Tc1::mScarlet-I and Tc1::sfgfp in somatic tissues ( Figure 3C ). Based upon these results and results presented below, F54D8.6 was named s uppressor of transposon m obilization ( stm ) -1 . Because stm-1(gg830) encodes a premature (Q10) stop in the predicted stm-1 open reading frame ( Figure 3A ), stm-1(gg830) is likely loss-of-function, indicating that a wild-type function of STM-1 is to limit Tc1 excision. stm-1 is predicted to encode a protein of unknown function, which lacks obvious homologs outside of Caenorhabditae . We asked if, as expected, HRPC-1 and STM-1 regulate endogenous Tc1 elements. We used qPCR to quantify rates of Tc1 excision from two of the ≅30 endogenous Tc1 elements present in the C. elegans N2 Bristol strain genome ( Figure 3D ). The analysis revealed a 3-8 fold increase in Tc1 excision events at these endogenous Tc1 loci in both hrpc-1 or stm-1 mutant animals, confirming that HRPC-1 and STM-1 are regulators of endogenous Tc1 elements ( Figure 3D ). The C. elegans soma and germline utilize different transposon regulatory strategies We used our reporter strains to assess the rate of germline Tc1 excision in hrpc-1 and stm-1 animals. Interestingly, hrpc-1 and stm-1 animals did not exhibit detectable increases in germline Tc1 excision from Tc1 reporter genes, as was apparent in somatic cells in these animals ( Table 2 ). The data suggests that loss of HRPC-1 or STM-1 does not increase Tc1 excision rates in the germline at least to levels detectable in these assays (see below). To test the idea that HRPC-1 and STM-1 act primarily in the soma to regulate Tc1 , we used an auxin-degron system to deplete a degron::GFP tagged HRPC-1 from either the germline or the soma. Fluorescent imaging confirmed auxin-dependent depletion of HRPC-1::AID::GFP from the soma or the germline (Figure S3). Depletion of HRPC-1 from somatic tissues resulted in an increase in Tc1 excision ( Figure 3E ). Note that in these experiments, animals that were not treated with auxin expressed less HRPC-1::AID::GFP (Figure S3), and showed Tc1 excision rates greater than control animals not expressing somatic TIR1 ( Figure 3E ), suggesting that some HRPC-1::AID::GFP depletion occurs even in the absence of auxin treatment. Depletion of HRPC-1::AID::GFP from the germline, however, did not impact Tc1 excision ( Figure 3E ). Similarly, ablation of germ cells, using a temperature sensitive glp-1(q224) allele in either hrpc-1 or stm-1 animals did not decrease endogenous Tc1 excision rates (Figure S4). These data show that loss of HRPC-1 results in an increase in Tc1 excision in the soma, but not in the germline, at least to levels detectable in these assays (see below). To assess if HRPC-1 might regulate Tc1 excision in the germline at levels below the detection threshold of our assays, we asked if loss of HRPC-1 would enhance germline Tc1 excision in animals lacking the germline RNAi factor RDE-3. To address this question, we assayed germline Tc1 excision frequencies in rde-3 and rde-3; hrpc-1 double mutant animals. The analysis revealed a >10 fold increase in the germline Tc1 excision rates in rde-3; hrpc-1 mutants compared to rde-3 single mutants for Tc1::sfgfp ( Table 2 ) and a 2x increase in excision of an endogenous Tc1 element (Figure S5). The data suggests that HRPC-1 acts in both the soma and germline to prevent Tc1 excision, however, the consequences of losing HRPC-1 in germ cells are minor in animals harboring an intact RNAi system. HRPC-1 and STM-1 are ubiquitously expressed nuclear proteins that interact in vivo To identify where and when hrpc-1 and stm-1 are expressed, we introduced GFP or mScarlet-I tags to the hrpc-1 and stm-1 genes, respectively. Wild-type animals develop from egg to larval stage four (L4) in ≅48 hours. hrpc-1 and stm-1 mutant animals exhibit delays in larval development; these animals took ≅72 and 60 hours respectively to reach the L4 stage of development (Figure S6). Epitope-tagged hrpc-1::gfp or stm-1::mScarlet-I expressing animals did not exhibit delays in larval development, indicating that the fluorescent tags introduced into these genes did not impact protein function, at least with regards to developmental timing (Figure S6). Fluorescent microscopy revealed that HRPC-1::GFP and STM-1::mScarlet-I were expressed in most or all cells of the soma ( Figure 4A ) and the germline ( Figure 4B ), and in these cells, HRPC-1::GFP and STM-1::mScarlet-I localized predominantly within nuclei ( Figures 4A-B ). We conclude that HRPC-1 and STM-1 are ubiquitously expressed nuclear proteins. The shared expression patterns of HRPC-1 and STM-1, and the shared developmental phenotypes of hrpc-1 and stm-1 mutants, hints that HRPC-1 and STM-1 might interact physically. To test this idea, we introduced hemagglutinin (HA) epitopes to the hrpc-1 gene for immunoprecipitation mass spectrometry (IP-MS) experiments. hrpc-1::3xha animals did not exhibit delays in larval development, suggesting the HA epitopes do not negatively impact HRPC-1 function (Figure S6). IP-MS of HRPC-1::3xHA identified STM-1 as the most enriched protein in HRPC-1::3xHA precipitates ( Fig. 4C-D , Figure S7, and Supplementary Table 1), suggesting that HRPC-1 and STM-1 associate physically in vivo . Download figure Open in new tab Figure 4. HRPC-1 and STM-1 localize to the nucleus and interact in vivo . (A and B) Representative confocal images showing expression of HRPC-1::GFP in the soma (A) and germline (B). (Top rows) Nuclei are marked with the nuclear pore protein TagRFP::NPP-9. (Bottom rows) STM-1::mScarlet-I colocalizes with HRPC-1::GFP in the nucleus. Scale bar, 40 μm ( A ), 5 μm ( B ). (C and D) IP-MS experiment to identify HRPC-1 interactors. ( C ) Gel image showing immunoblotting of HA and GAPDH in indicated samples from the IP-MS experiment. Wild-type strain is the control. FT, flow-through fraction. ( D ) Number of unique and total peptides of the top 5 proteins that were only identified from hrpc-1::3xha IP-MS. Results of another independent replicate are shown in Figure S7. In the independent IP-MS replicate, STM-1 was the most common co-precipitating protein and the other 3 proteins did not reproduce. WT, wild-type. HRPC-1/STM-1 associate with Tc1 -containing RNAs Because HRPC-1 possesses an RRM RNA binding domain, we wondered if HRPC-1 might interact with Tc1 RNA. To test this idea, we used hrpc-1::3xha animals to conduct anti-HA immunoprecipitation (IP) and then RT-qPCR to quantify co-precipitating RNAs. The analysis showed that IP of HRPC-1::3xHA enriched Tc1 RNA in the precipitate >300x over levels of Tc1 RNA observed in control IPs, which employed animals not expressing HRPC-1::3xHA ( Figures 5A-B ). HRPC-1::3xHA IP did not enrich a randomly selected mRNA ( cdc-42 ), which does not contain a Tc1 element ( Figure 5B ), suggesting that the interaction of HRPC-1 with the Tc1 RNA is at least somewhat specific. Introduction of the A132V RRM variant into HRPC-1::3xHA by CRISPR negatively impacted the expression of HRPC-1::3xHA (see Figure 5A -input, S8A-input, Figure S8C). Nonetheless, when levels of Tc1 RNA in the pellet were normalized to levels of immunoprecipitated HRPC-1 protein, HRPC-1(A132V)::3xHA failed to enrich Tc1 RNA, suggesting that the A132C RRM mutation disrupts HRPC-1/ Tc1 RNA interactions ( Fig. 5B and S8B). We next asked if STM-1 might be required for HRPC-1::3xHA to associate with Tc1 -containing RNAs. Again, HRPC-1 was poorly expressed in stm-1 mutant backgrounds (see Figure 5A -input, S8A-input). Nonetheless, when levels of RNA in the pellet were normalized to levels of immunoprecipitated HRPC-1 protein, HRPC-1::3xHA co-precipitated with less Tc1 RNA in an stm-1 mutant than in stm-1(+) animals, although HRPC-1::3xHA retained some ability to associate with Tc1 in the absence of STM-1 ( Fig. 5B and S8B). The data suggest that HRPC-1 associates with Tc1 -encoded RNA and that this association depends, at least in part, on STM-1 and the RRM domain of HRPC-1. The data also support the idea that HRPC-1 and STM-1 act together in a pathway to repress Tc1 . Download figure Open in new tab Figure 5. HRPC-1 binds Tc1 RNA and suppresses Tc1 RNA levels. (A and B) RNA immunoprecipitation experiment for detection of HRPC-1 interaction with Tc1 RNA. (A) Representative image of HA immunoblotting of indicated proteins (tubulin, loading control). FT, flow-through fraction. IP, immunoprecipitation fraction. WT, wild-type. (B) RT-qPCR quantification of indicated RNAs in HRPC-1::HA immunoprecipitates. Levels of RNA in immunoprecipitates were normalized to levels of RNA in inputs. Mean enrichment level of wild-type sample was set as 1. ns: not significant; ***, p ≤ 0.001 (Two-way ANOVA with Šídák’s multiple comparisons test.) Data are from 3 biological replicates. Mean ± SD. (C) (Left) Primers for RT-qPCR of total Tc1 (includes unspliced and spliced Tc1 ) or spliced Tc1 RNA as indicated. The forward primer for detecting spliced Tc1 RNA spans the Tc1 exon-exon junction. (Right) RT-qPCR quantification of total Tc1 and spliced Tc1 RNA levels in hrpc-1 and stm-1 . **, p ≤ 0.01; ***, p ≤ 0.001 (one-way ANOVA with Dunnett’s multiple comparisons test.). Data are from 3 biological replicates. Mean ± SD. WT, wild-type. HRPC-1/STM-1 regulate Tc1 -containing RNAs Since HRPC-1 associates with Tc1 RNA, we wondered whether HRPC-1/STM-1 might regulate Tc1 RNA abundance and/or processing. Indeed, we observed elevated (5-10x) levels of Tc1 RNA in hrpc-1 and stm-1 mutant animals ( Figure 5C , total Tc1 ), indicating that HRPC-1/STM-1 negatively regulates Tc1 RNA. While conducting the above RT-PCR analysis, we noticed that the size of Tc1 RNA differed in hrpc-1 and stm-1 mutants: In wild type animals, RT-PCR of Tc1 generated a single PCR product while in hrpc-1 and stm-1 mutants the RT-PCR produced two bands, which differed in length by ∼40 base pairs ( Figure 6 ). The Tc1 RNA contains a single 41 nucleotide intron that must be spliced for the Tc1 mRNA to be translated [ 52 ]. For unknown reasons (see discussion), most Tc1 RNA is not spliced in wild-type C. elegans [ 53 ]. Sequencing revealed that the two PCR products observed in hrpc-1 and stm-1 mutants were spliced and unspliced Tc1 RNAs (Figure S9). RT-qPCR analysis confirmed that Tc1 RNA is more efficiently spliced in the absence of HRPC-1 and STM-1 ( Figure 5C , spliced Tc1 ). These results suggest that HRPC-1 and STM-1 could suppress Tc1 RNA levels- and Tc1 activity-by inhibiting intron processing of Tc1 , thereby decreasing Tc1 RNA stability. Download figure Open in new tab Figure 6. HRPC-1 and STM-1 affect splicing of Tc1 RNAs. Electrophoresis of RT-PCR products showing Tc1 intron splicing efficiency in animals of the indicated genotypes. Triangles indicate bands for unspliced (filled) and spliced (hollow) RNAs. Splicing of the Tc1 intron is increased in hrpc-1 and stm-1 mutants compared to WT. Splicing of a control pmp-3 intron is similarly efficient across all three genotypes. WT, wild type. In summary, we report the development of fluorescence-based reporter genes for studying transposon regulation in a living animal. Using these tools, we show that C. elegans uses different systems to regulate Tc1 excision in its germline and soma and we identify a pathway involving HRPC-1 and STM-1 that limits Tc1 excision primarily in the soma, likely through binding Tc1 RNA to limit Tc1 intron splicing. Discussion Here, we develop fluorescence-based reporter genes to visualize transposon excision at single-cell level resolution in a living animal. We use these reporter genes to show that C. elegans employs different strategies to regulate Tc1 in its soma and germline. Consistent with previous research [ 54 ], we find that cytoplasmic factors that localize to- and/or help organize-germ granules (DEPS-1 and ZNFX-1) [ 46 , 47 ], and a nuclear factor that acts in the nuclear branch of the RNAi pathway (NRDE-2) [ 51 ], contribute to transposon suppression in the germline. The results reinforce the idea that the germline anti-transposon system is complex and multi-faceted, involving both cytoplasmic and nuclear branches. Our results also show that, surprisingly, the germline RNAi pathway is not employed in the soma to regulate Tc1 , despite many of the germline RNAi components being expressed in the soma, where they contribute to RNAi-based gene silencing [ 40 , 43 , 55 ]. Why wouldn’t C. elegans use the germline RNAi system to control transposons, such as Tc1 , in somatic tissues? The RNAi pathway is complex, employing dozens of proteins subcompartmentalized into both membrane and non-membrane delimited organelles in germ cells [ 56 ]. It is possible that C. elegans , and perhaps other organisms, choose not to deploy RNAi to combat transposons in their somatic tissues because the fitness costs of somatic transposition are not as high in the soma as they are in the germline and because the system is too energetically costly to merit deployment. Rather, we find that C. elegans uses an HNRNPC/STM-1-based system to regulate Tc1 in its soma. Interestingly, human HNRNPC binds and prevents exonization of Alu retrotransposons, which are the most abundant transposons in the human genome [ 26 , 27 ]. Thus, the role of HRPC-1 in regulating transposon-encoded RNAs may be conserved. Future studies are needed to assess if the role of HNRNPC in regulating transposons in nematodes and mammals reflects an ancient function of the HNRNPC proteins, or if the shared functions are due to convergent evolution. It will also be of interest to determine the breadth of transposon families regulated by HNRNPC-like proteins in eukaryotes. Human HNRNPC is thought to bind antisense-oriented Alu RNAs, at least in part, by recognizing single-stranded poly-U tracts present in Alu RNAs synthesized from genes containing antisense, intronic Alu insertions [ 27 ]. By recognizing poly-U tracts, HNRNPC prevents binding of the splicing factor U2AF65, thereby preventing Alu elements from becoming exonized into host mRNAs [ 27 ]. It is possible that C. elegans HRPC-1 may detect the Tc1 RNA by binding poly-U tracts-the Tc1 RNA contains 19 poly-U tracts ≥5 base pairs. Additionally, RNA secondary structure can influence HNRNPC interaction with RNA [ 57 , 58 ] and transposon-encoded RNAs often adopt secondary structures [ 26 , 41 ], which may also serve as a recognition feature for HRPC-1/HNRNPC. Understanding how HRPC-1 and HNRNPC identify transposon-encoded RNAs for neutralization will be an important question for future studies to address. We find that splicing of the Tc1 intron is more efficient in hrpc-1 and stm-1 mutant animals, which would be expected to increase transposase expression and, therefore, Tc1 mobilization. Intriguingly, splicing of the third intron of the DNA transposon P element in Drosophila is also actively regulated, and this splicing determines whether the translated protein is active as a transposase [ 59 ]. In Drosophila somatic cells, splicing of P elements is repressed by RNA-binding proteins that recognize sequence specific elements upstream of the alternatively spliced third intron [ 60 ]. Taken together with our observations, the data hint that the regulation of transposon splicing could be a common strategy employed by animal cells to regulate DNA transposons in the soma. A final question emerging from our work is why would C. elegans Tc1 and human Alu possess signals that allow HRPC-1/HNRNPC to bind and regulate their splicing. Transposons cannot survive without their hosts, and transposition in the soma does not benefit the transposon. Therefore, we speculate that transposons are under selective pressure to evolve sequences that allow HRPC-1/HNRNPC (and perhaps other systems) to limit their splicing in the soma in order to limit transposon exonization into host genes, thereby minimizing the fitness costs incurred by both host and parasite. Asking if C. elegans HRPC-1 prevents the exonization of Tc1 , or other transposons, into host mRNAs will be a strong test of this latter idea. Materials and Methods Strains All strains were cultivated on standard Nematode Growth Medium (NGM) plates and maintained at 20°C with Escherichia coli strain OP50 as food source unless otherwise noted [ 62 ]. glp-1 ( q224ts ) animals were maintained at 15°C. CRISPR strains were made with the CRISPR/Cas9 strategies described by Arribere et al., or Ghanta and Mello [ 63 , 64 ]. Guide RNAs were chosen according to the CRISPOR.org tool [ 65 ]. Tc1::mScarlet-I and Tc1::sfgfp reporter strains were made via CRISPR/Cas9 with Saptrap [ 66 ] combined with self-excising cassette selection strategy [ 66 , 67 ]. List of strains and details is available in Supplementary Table 2. Tc1 reporter plasmids construction Tc1::mScarlet-I and Tc1::sfgfp reporter plasmids for CRISPR/Cas9 injection to make Tc1 reporter strains were constructed by using Saptrap to assemble components into the pDD379 vector [ 66 ]. Tc1::mScarlet-I reporter plasmid donor sequence consists of Tc1 (1613 base pairs, including a TA dinucleotide at the 3’ end) inserted after the TA dinucleotide at position -7 to -8 of the 622 base pair eft-3 promoter, followed by mScarlet-I sequence with nuclear-localization sequences (NLS) (SV40 NLS at 5’ end; egl-13 NLS at 3’ end) and the tbb-2 3’UTR. Tc1::sfgfp reporter plasmid donor sequence consists of a 523 base pair rpl-28 promoter, followed by a start codon and an 18 base pair flexible linker before the Tc1 sequence (1615 base pairs, including TA dinucleotide at both 5’ and 3’ ends). The Tc1 sequence is followed by another 18 base pair flexible linker before the sfgfp sequence with NLSs (SV40 NLS at 5’ end; egl-13 NLS at 3’ end of sfgfp ) and the unc-54 3’UTR. The nuclear localized sfgfp sequence lacks a start codon. The Tc1 sequence in Tc1::mScarlet-I and Tc1::sfgfp plasmid constructs is the sequence of Tc1 with WormBase ID WBTransposon00000015, which does not contain any polymorphisms [ 11 ]. Tc1::mScarlet-I reporter plasmid guide RNA and homology arm components were designed to insert the reporter at Chr I: -5.32 cM. Tc1::sfgfp reporter plasmid guide RNA and homology arm components were designed to insert the reporter at Chr III: -0.85 cM. These sites were chosen according to insertion sites established for Mos1 mediated single copy insertion (MosSCI) that allow germline expression of inserts [ 68 , 69 ]. Microscopy In Figures 1B , 1C , 1E images, animals were immobilized with 0.05% sodium azide and mounted on glass slides; in Fig S3 images, animals were mounted on glass slides without immobilization. Images in Figures 1B , 1C , 1E , S3 were taken using the Axio Observer.Z1 fluorescent microscope (Zeiss) with the Plan-Apochromat 20x/0.8 M27 objective or the Plan-Apochromat 63x/1.4 Oil DIC M27 objective. Images were acquired using ORCA-Flash4.0 V2 digital camera (Hamamatsu) and ZEN 2 (blue edition) software (Zeiss). For Figures 3C , S2, and 4A-B images, larval stage 4 (L4) animals ( Figures 3C and S2) or one day old adult animals ( Figures 4A-B ) were immobilized with polystyrene beads (Polysciences, Cat #00876) and mounted on 10% agarose pads. Images were taken using the Dragonfly 505 spinning-disk confocal microscope with iXon Ultra 888 EMCCD camera (Andor Technologies) using Nikon 60x/1.40 ( Figure 3C , S2, 4A) or 100x/1.45 ( Figure 4B ) oil immersion objectives. Images were processed using Fiji [ 70 ]. Microscope images in Figures 1B , 1C, 3C and S2 were stitched together from two or more images because a single image was not able to capture the full x-y dimension. Figure 3C and S2 images are maximum intensity projections of z-stack images taken at a step size of 0.5 μm spanning the thickness of the animal. Tc1 excision footprint analysis Tc1::mScarlet-I ; glp-1 ( q224ts ) strains were bleach-synchronized and grown from embryonic stage at 25°C for 3 days to reach adult stage. Adult worms were washed off the plate with M9 buffer and lysed with worm lysis buffer (50mM KCl, 10mM pH8.3 Tris, 2.5mM MgCl 2 ) with proteinase K (0.2 mg/ml). Lysates were used as templates for PCR reactions using primers: F: 5’-CTACCGTCCGCACTCTTC-3’; R: 5’-CTTACGCTTCTTCTTTGGC-3’. PCR products were run on an agarose gel, and band size around 100 base pairs was excised and gel purified (Qiagen #27206). Purified PCR products were TA-cloned using the pGEM-T Easy Vector System (Promega, A1360), according to the manual instructions. White transformants were isolated, grown in liquid culture, and miniprepped. Plasmids were sent for Sanger sequencing using the primer M13F-40: 5’-GTT TTC CCA GTC ACG AC-3’. Soma Tc1 excision rate quantification For the soma Tc1 excision assay in Table 1 , for all genotypes except hrpc-1 and stm-1 , L4 animals (P0) were singled onto 6 cm NGM plates and kept at 20°C for their progeny animals to grow. 4-5 days later, each plate was examined under the AxioZoom.v16 fluorescence microscope (Zeiss) with the ApoZ 1.5x/0.37 objective. Progeny animals (F1) of the original singled animal were visually scored for presence or absence of somatic cells with mScarlet-I expression, to determine the number of progeny animals on a single plate that had somatic mScarlet-I expression. All visual scoring of soma mScarlet-I expression was done with the examiner blinded to genotype. This data is also shown as Figure 2D . For hrpc-1 and stm-1 , percentage of animals showing somatic cell mScarlet-I (+) was 100%, and L4 stage animals were examined under the Dragonfly 505 spinning-disk confocal microscope using the Nikon 60x/1.40 oil immersion objective to quantify number of somatic cells with mScarlet-I expression per animal. This data is also shown as Figure 3B . Germline Tc1 excision rate quantification Previous labs have established a Poisson distribution method for estimating germline transposon excision rate in C. elegans [ 19 , 36 , 37 , 71 ]. This method relies on establishing single lineages of animals, which are then scored as a population for whether or not excision event(s) have occurred at all in the span of the propagation time. Taking into account the amount of animals present in each lineage at time of scoring and the proportion of lineages that have excision event(s), a germline excision rate is calculated using the Poisson distribution formula [ 19 , 36 ]: f = - (ln(N/T))/n, f: germline Tc1 excision frequency, T: total number of lineages scored, N: total number of lineages without germline Tc1 excision events, n: average number of animals per lineage. This calculation method addresses the concern of counting a single germline Tc1 excision event multiple times, since a single animal that undergoes a germline excision event will generate many progeny presenting with germline excision events that result from inheriting the excised Tc1 locus instead of undergoing actual independent germline excision events (jackpot effect) [ 19 , 36 , 37 , 71 ]. In our germline excision experiments ( Table 2 ), L4 animals were singled onto 6 cm NGM plates and allowed to propagate at 20°C. 20x concentrated OP50 was added to plates during the interval of growth. After several days of propagation, when plates have reached either F1 or F2/3 generations, each plate (i.e. each lineage) was scored for presence or absence of germline Tc1 excision event(s). Scoring was done under the AxioZoom.v16 fluorescence microscope (Zeiss) with the ApoZ 1.5x/0.37 objective for Tc1::mScarlet-I background animals and under the Leica M165 FC with a Plan APO 1.0x objective for Tc1::sfgfp background animals. A germline excision event is indicated by presence of an animal in which all cells are expressing mScarlet-I or sfGFP. The generation at which animals are scored for Tc1 excision (F1 or F2/3) varies with genotype and depends on how frequent germline Tc1 excision is in that particular genotype. The number of animals on each plate at time of scoring was estimated by counting animals in one region of the plate, then multiplying by plate area. The average number of animals on each plate (“n” variable in the Poisson distribution formula) is calculated by summing the number of animals on each plate for a genotype, then dividing this sum by the total number of plates for that genotype. Germline Tc1 excision rates were calculated using the Poisson distribution formula. In genotypes where no germline Tc1 excision event was detected, germline Tc1 excision rates were indicated as less than the rate that would have been if one plate of all plates scored had germline Tc1 excision event(s). All visual scoring was done with the examiner blinded to genotype. Forward genetic screen for Tc1 excision regulators Tc1::mScarlet-I; Tc1::sfgfp strains were treated with 47 mM ethyl methanesulfonate for four hours at 23°C (P0 generation), then washed with M9 buffer. At the F2, F3, F4 generations, mutants were visually screened under the AxioZoom.v16 fluorescence microscope (Zeiss) and mutants that had a high number of somatic cells expressing mScarlet-I were singled onto new plates. After the singled mutants gave rise to progeny, their progeny animals were verified to have high levels of somatic mScarlet-I and somatic sfGFP using the Axio Observer.Z1 fluorescent microscope (Zeiss). The screen was done in two rounds; for the first round, ∼69,000 haploid genomes were screened, and mutants were picked at F2 and F3 generations; for the second round, ∼180,000 haploid genomes were screened, and mutants were picked at F2, F3 and F4 generations. Mutant genomic DNA was extracted and sent for whole genome sequencing (Biopolymers Facility, Harvard Medical School). Based on whole genome sequencing and positional mapping information, causal alleles were identified. Tc1 excision qPCR assay L4 stage animals were lysed in worm lysis buffer (50mM KCl, 10mM pH8.3 Tris, 2.5mM MgCl 2 ) with proteinase K (0.2 mg/ml). DNA lysates were used for qPCR assays using the iTaq Universal SYBR Green Supermix (Bio-Rad, 1725121), as instructed by the manual, and run on CFX Connect Real-Time PCR Detection System (Bio-Rad, 1855201). qPCR conditions were as follows: Initial denaturation at 95°C for 5 min followed by 40 cycles of denaturation at 95°C for 15 sec and annealing/extension/plate read at 61°C for 30 sec. Tc1 excision fold changes were calculated using the comparative Cq method [ 72 ]. Mean Cq value of wild-type strain was used as the baseline for calculating Tc1 excision fold change for each experiment. eft-3 was used as the housekeeping gene. Primers used for qPCR analysis to detect Tc1#20 and Tc1#33 excision are listed in Supplementary Table 1. Multiple sequence alignment Multiple sequence alignment was performed using MUSCLE v3.8.31 in Jalview v2.11.4.1 [ 73 – 75 ]. Protein sequences used for alignment were imported from Uniprot [ 76 ]. Developmental timing assay Ten gravid adult animals were placed on a plate, allowed to lay eggs for 3 hours, then removed. 51 hours post egg lay, plates were scored for percentage of animals that reached L4 stage. At least 15 animals were scored for each plate. Visual scoring was done with the examiner blinded to genotype. Auxin-induced depletion Synchronized L1 animals were placed on seeded NGM plates with or without 1 mM auxin for 48 hours at 20°C. L4 animals were harvested for imaging or lysed for qPCR analysis. Immunoprecipitation-mass spectrometry (IP-MS) Approximately 10 5 synchronized adult animals (grown for 68-72 hours from L1 stage at 20°C) were used in each sample. Animals were harvested and washed three times with 10 mL M9 buffer, then flash frozen in liquid nitrogen and stored at -80°C until use. To prepare worm lysates, frozen worm beads were ground into fine powder with liquid nitrogen-cooled mortar and pestle. The worm powder was transferred into 10 mL of lysis buffer (10 mM HEPES-NaOH (pH 7.5), 50 mM NaCl, 2.5 mM MgCl 2 , 1 mM EDTA, 10% Glycerol, 0.5 mM DTT, 1 mM phenylmethylsulfonyl fluoride, 0.25% Triton X-100) supplemented with 1x cOmplete protease inhibitor cocktail (Roche, 11697498001). After rotating at 4°C for 15 minutes, the lysate was briefly centrifuged and then passed through a 0.45 µm syringe filter (Millipore, SLHPR33RS). Lysate was then incubated with either anti-HA agarose beads (Millipore, A2095) or anti-HA magnetic beads (Thermo Scientific, 88836) at 4°C for 3-4 hours and washed four times with 1mL lysis buffer. To elute bound proteins, beads were incubated with 0.5 M NH 4 OH at 37°C for 20 minutes. The eluate was vacuum dried and resuspended in 100 µl 10 mM Tris-HCl (pH 8.5) and sent to The Taplin Biological Mass Spectrometry Facility at Harvard Medical School for analysis. RNA immunoprecipitation (RIP) Approximately 10,000 synchronized L4 animals of each genotype were flash frozen in liquid nitrogen. Worm pellets were ground to powder with liquid nitrogen-cooled mortar and pestle, and resuspended with lysis buffer (20 mM Tris-HCl, pH7.5, 200 mM NaCl, 2.5 mM MgCl 2 , 10% glycerol, and 0.5% IGEPAL CA-630 supplemented with 1x cOmplete protease inhibitor cocktail (Roche, 11697498001) and 80 U/ml RNAseOUT (Invitrogen, 10777019)). The lysate was passed through a 0.45 μm syringe filter (Millipore, SLHPR33RS), then incubated with anti-HA magnetic beads (Thermo Scientific, 88836) at 4°C for 2 hours. After washing three times with 1 ml lysis buffer, 10% of the beads was saved for immunoblotting. TRIzol (Invitrogen, 15596026) was added directly to the rest of the beads for RNA extraction. Primers used for Tc1 RNA and cdc-42 mRNA detection are listed in Supplementary Table 1. RT-qPCR for spliced/total Tc1 RNA levels and RT-PCR to detect Tc1 RNA splicing Synchronized animals were frozen in TRIzol and total RNA was extracted using ZYMO Direct-zol RNA extraction kit with on-column DNase I digestion. cDNA synthesis was performed using SuperScript™ III First-Strand Synthesis System (Invitrogen, 18080051) with random hexamers following manufacturer’s protocols. Threshold cycle (Ct) numbers were obtained using iTaq Universal SYBR Green Supermix (Bio-Rad, 1725120) on Bio-Rad CFX Duet Real-Time PCR system (Bio-Rad, 12016265). For RT-PCR to check intron splicing efficiency, PCR products were resolved on 2-4% agarose gels. Primers used are listed in Supplementary Table 1. Author Contributions Conceptualization: C.C., S.K,; Investigation: C.C., D.C., D.J.P; Methodology: C.C., D.C.; Formal analysis: C.C., D.C.; Writing: C.C., S.K.; Supervision: S.K.; Funding acquisition: S.K. Data Availability Strains and plasmids are available upon request. Strains used in this study are listed and described in Supplementary Table 2. Complete data of HRPC-1::3xHA IP-MS (two replicates) is available in Supplementary Table 1. Study Funding This work was supported by NIH grant R35GM148206 (S.K.) In May of 2025, R35GM148206 was terminated by the US government. S.K. thanks Harvard University, Harvard Medical School, and the Department of Genetics at Harvard Medical School for support, following grant termination. Supplemental Information Figures S1-S9. Supplementary Tables 1 -2 . Acknowledgements We thank past and previous members of the Kennedy lab for helpful discussions and comments. We thank Dr. David Lowe and Dr. Gang Wan for help on mutant mapping genome analysis. We thank the BPF Genomics Core Facility at Harvard Medical School for their expertise and instrument availability that supported this work. Funder Information Declared National Institute of Health , R35GM148206 Harvard Medical School, https://ror.org/03wevmz92 Harvard University, https://ror.org/03vek6s52 References 1. ↵ Huang CRL , Burns KH , Boeke JD . Active transposition in genomes . Annu Rev Genet . 2012 ; 46 : 651 – 75 . OpenUrl CrossRef PubMed Web of Science 2. ↵ Klein SJ , O’Neill RJ . Transposable elements: genome innovation, chromosome diversity, and centromere conflict . Chromosome Res . 2018 ; 26 : 5 – 23 . OpenUrl CrossRef PubMed 3. ↵ Slotkin RK , Martienssen R . Transposable elements and the epigenetic regulation of the genome . Nat Rev Genet . 2007 ; 8 : 272 – 85 . OpenUrl CrossRef PubMed Web of Science 4. Levin HL , Moran JV . Dynamic interactions between transposable elements and their hosts . Nat Rev Genet . 2011 ; 12 : 615 – 27 . OpenUrl CrossRef PubMed 5. Wolf G , Greenberg D , Macfarlan TS . Spotting the enemy within: Targeted silencing of foreign DNA in mammalian genomes by the Krüppel-associated box zinc finger protein family . Mob DNA . 2015 ; 6 : 17 . OpenUrl CrossRef PubMed 6. ↵ Deniz Ö , Frost JM , Branco MR . Regulation of transposable elements by DNA modifications . Nat Rev Genet . 2019 ; 20 : 417 – 31 . OpenUrl CrossRef PubMed 7. ↵ Cordaux R , Batzer MA . The impact of retrotransposons on human genome evolution . Nat Rev Genet . 2009 ; 10 : 691 – 703 . OpenUrl CrossRef PubMed Web of Science 8. ↵ Feschotte C , Pritham EJ . DNA transposons and the evolution of eukaryotic genomes . Annu Rev Genet . 2007 ; 41 : 331 – 68 . OpenUrl CrossRef PubMed Web of Science 9. ↵ Fischer SEJ . Activity and Silencing of Transposable Elements in C. elegans . DNA . 2024 ; 4 : 129 – 40 . OpenUrl CrossRef 10. ↵ Plasterk RH . The Tc1/mariner transposon family . Curr Top Microbiol Immunol . 1996 ; 204 : 125 – 43 . OpenUrl PubMed Web of Science 11. ↵ Fischer SEJ , Wienholds E , Plasterk RHA . Continuous exchange of sequence information between dispersed Tc1 transposons in the Caenorhabditis elegans genome . Genetics . 2003 ; 164 : 127 – 34 . OpenUrl Abstract / FREE Full Text 12. ↵ Haig D . Transposable elements: Self-seekers of the germline, team-players of the soma . Bioessays . 2016 ; 38 : 1158 – 66 . OpenUrl CrossRef PubMed 13. ↵ Lee E , Iskow R , Yang L , Gokcumen O , Haseley P , Luquette LJ 3rd , et al. Landscape of somatic retrotransposition in human cancers. Science . 2012 ; 337 : 967 – 71 . OpenUrl 14. Scott EC , Gardner EJ , Masood A , Chuang NT , Vertino PM , Devine SE . A hot L1 retrotransposon evades somatic repression and initiates human colorectal cancer . Genome Res . 2016 ; 26 : 745 – 55 . OpenUrl Abstract / FREE Full Text 15. Rodić N , Sharma R , Sharma R , Zampella J , Dai L , Taylor MS , et al. Long interspersed element-1 protein expression is a hallmark of many human cancers . Am J Pathol . 2014 ; 184 : 1280 – 6 . OpenUrl CrossRef PubMed 16. ↵ Payer LM , Burns KH . Transposable elements in human genetic disease . Nat Rev Genet . 2019 ; 20 : 760 – 72 . OpenUrl CrossRef PubMed 17. ↵ Teixeira FK , Okuniewska M , Malone CD , Coux R-X , Rio DC , Lehmann R . piRNA-mediated regulation of transposon alternative splicing in the soma and germ line [Internet] . Nature . 2017 . p. 268 – 72 . Available from : doi: 10.1038/nature25018 OpenUrl CrossRef PubMed 18. ↵ Pélisson A , Song SU , Prud’homme N , Smith PA , Bucheton A , Corces VG . Gypsy transposition correlates with the production of a retroviral envelope-like protein under the tissue-specific control of the Drosophila flamenco gene . EMBO J . 1994 ; 13 : 4401 – 11 . OpenUrl CrossRef PubMed Web of Science 19. ↵ Eide D , Anderson P . Insertion and excision of Caenorhabditis elegans transposable element Tc1 . Mol Cell Biol . 1988 ; 8 : 737 – 46 . OpenUrl Abstract / FREE Full Text 20. Eide D , Anderson P . Transposition of Tc1 in the nematode Caenorhabditis elegans . Proc Natl Acad Sci U S A . 1985 ; 82 : 1756 – 60 . OpenUrl Abstract / FREE Full Text 21. ↵ Collins J , Saari B , Anderson P . Activation of a transposable element in the germ line but not the soma of Caenorhabditis elegans . Nature . 1987 ; 328 : 726 – 8 . OpenUrl CrossRef PubMed 22. ↵ Beyer AL , Christensen ME , Walker BW , LeStourgeon WM . Identification and characterization of the packaging proteins of core 40S hnRNP particles . Cell . 1977 ; 11 : 127 – 38 . OpenUrl CrossRef PubMed Web of Science 23. ↵ König J , Zarnack K , Rot G , Curk T , Kayikci M , Zupan B , et al. iCLIP reveals the function of hnRNP particles in splicing at individual nucleotide resolution . Nat Struct Mol Biol . 2010 ; 17 : 909 – 15 . OpenUrl CrossRef PubMed Web of Science 24. Fischl H , Neve J , Wang Z , Patel R , Louey A , Tian B , et al. hnRNPC regulates cancer-specific alternative cleavage and polyadenylation profiles . Nucleic Acids Res . 2019 ; 47 : 7580 – 91 . OpenUrl CrossRef PubMed 25. ↵ Gruber AJ , Schmidt R , Gruber AR , Martin G , Ghosh S , Belmadani M , et al. A comprehensive analysis of 3’ end sequencing data sets reveals novel polyadenylation signals and the repressive role of heterogeneous ribonucleoprotein C on cleavage and polyadenylation . Genome Res . 2016 ; 26 : 1145 – 59 . OpenUrl Abstract / FREE Full Text 26. ↵ Deininger P . Alu elements: know the SINEs . Genome Biol . 2011 ; 12 : 236 . OpenUrl CrossRef PubMed 27. ↵ Zarnack K , König J , Tajnik M , Martincorena I , Eustermann S , Stévant I , et al. Direct competition between hnRNP C and U2AF65 protects the transcriptome from the exonization of Alu elements . Cell . 2013 ; 152 : 453 – 66 . OpenUrl CrossRef PubMed Web of Science 28. Wilusz J , Shenk T . A uridylate tract mediates efficient heterogeneous nuclear ribonucleoprotein C protein-RNA cross-linking and functionally substitutes for the downstream element of the polyadenylation signal . Mol Cell Biol . 1990 ; 10 : 6397 – 407 . OpenUrl Abstract / FREE Full Text 29. Kim JH , Paek KY , Choi K , Kim T-D , Hahm B , Kim K-T , et al. Heterogeneous nuclear ribonucleoprotein C modulates translation of c-myc mRNA in a cell cycle phase-dependent manner . Mol Cell Biol . 2003 ; 23 : 708 – 20 . OpenUrl Abstract / FREE Full Text 30. Shetty S . Regulation of urokinase receptor mRNA stability by hnRNP C in lung epithelial cells . Mol Cell Biochem . 2005 ; 272 : 107 – 18 . OpenUrl CrossRef PubMed Web of Science 31. Ray D , Kazan H , Cook KB , Weirauch MT , Najafabadi HS , Li X , et al. A compendium of RNA-binding motifs for decoding gene regulation . Nature . 2013 ; 499 : 172 – 7 . OpenUrl CrossRef PubMed Web of Science 32. Cieniková Z , Damberger FF , Hall J , Allain FH-T , Maris C . Structural and mechanistic insights into poly(uridine) tract recognition by the hnRNP C RNA recognition motif . J Am Chem Soc . 2014 ; 136 : 14536 – 44 . OpenUrl CrossRef PubMed 33. ↵ Swanson MS , Dreyfuss G . Classification and purification of proteins of heterogeneous nuclear ribonucleoprotein particles by RNA-binding specificities . Mol Cell Biol . 1988 ; 8 : 2237 – 41 . OpenUrl Abstract / FREE Full Text 34. ↵ Harris LJ , Rose AM . Somatic excision of transposable element Tc1 from the Bristol genome of Caenorhabditis elegans . Mol Cell Biol . 1986 ; 6 : 1782 – 6 . OpenUrl Abstract / FREE Full Text 35. ↵ Eide D , Anderson P . Transposition of Tc1 in the nematode Caenorhabditis elegans . Proc Natl Acad Sci U S A . 1985 ; 82 : 1756 – 60 . OpenUrl Abstract / FREE Full Text 36. ↵ Collins J , Saari B , Anderson P . Activation of a transposable element in the germ line but not the soma of Caenorhabditis elegans . Nature . 1987 ; 328 : 726 – 8 . OpenUrl CrossRef PubMed 37. ↵ Moerman DG , Waterston RH . Spontaneous unstable unc-22 IV mutations in C. elegans var. Bergerac . Genetics . 1984 ; 108 : 859 – 77 . OpenUrl Abstract / FREE Full Text 38. ↵ Vastenhouw NL , Fischer SEJ , Robert VJP , Thijssen KL , Fraser AG , Kamath RS , et al. A genome-wide screen identifies 27 genes involved in transposon silencing in C. elegans . Curr Biol . 2003 ; 13 : 1311 – 6 . OpenUrl CrossRef PubMed Web of Science 39. ↵ Preston MA , Porter DF , Chen F , Buter N , Lapointe CP , Keles S , et al. Unbiased screen of RNA tailing activities reveals a poly(UG) polymerase . Nat Methods . 2019 ; 16 : 437 – 45 . OpenUrl CrossRef PubMed 40. ↵ Shukla A , Yan J , Pagano DJ , Dodson AE , Fei Y , Gorham J , et al. poly(UG)-tailed RNAs in genome protection and epigenetic inheritance . Nature . 2020 ; 582 : 283 – 8 . OpenUrl CrossRef PubMed 41. ↵ Sijen T , Plasterk RHA . Transposon silencing in the Caenorhabditis elegans germ line by natural RNAi . Nature . 2003 ; 426 : 310 – 4 . OpenUrl CrossRef PubMed Web of Science 42. ↵ Chen C-CG , Simard MJ , Tabara H , Brownell DR , McCollough JA , Mello CC . A member of the polymerase beta nucleotidyltransferase superfamily is required for RNA interference in C. elegans . Curr Biol . 2005 ; 15 : 378 – 83 . OpenUrl CrossRef PubMed Web of Science 43. ↵ Zhang C , Montgomery TA , Gabel HW , Fischer SEJ , Phillips CM , Fahlgren N , et al. mut-16 and other mutator class genes modulate 22G and 26G siRNA pathways in Caenorhabditis elegans . Proc Natl Acad Sci U S A . 2011 ; 108 : 1201 – 8 . OpenUrl Abstract / FREE Full Text 44. ↵ Phillips CM , Montgomery TA , Breen PC , Ruvkun G . MUT-16 promotes formation of perinuclear mutator foci required for RNA silencing in the C. elegans germline . Genes Dev . 2012 ; 26 : 1433 – 44 . OpenUrl Abstract / FREE Full Text 45. ↵ Dodson AE , Kennedy S . Phase Separation in Germ Cells and Development . Dev Cell . 2020 ; 55 : 4 – 17 . OpenUrl CrossRef PubMed 46. ↵ Spike CA , Bader J , Reinke V , Strome S . DEPS-1 promotes P-granule assembly and RNA interference in C. elegans germ cells . Development . 2008 ; 135 : 983 – 93 . OpenUrl Abstract / FREE Full Text 47. ↵ Wan G , Fields BD , Spracklin G , Shukla A , Phillips CM , Kennedy S . Spatiotemporal regulation of liquid-like condensates in epigenetic inheritance . Nature . 2018 ; 557 : 679 – 83 . OpenUrl CrossRef PubMed 48. ↵ Dodson AE , Kennedy S . Phase Separation in Germ Cells and Development . Dev Cell . 2020 ; 55 : 4 – 17 . OpenUrl CrossRef PubMed 49. ↵ Guang S , Bochner AF , Pavelec DM , Burkhart KB , Harding S , Lachowiec J , et al. An Argonaute transports siRNAs from the cytoplasm to the nucleus . Science . 2008 ; 321 : 537 – 41 . OpenUrl Abstract / FREE Full Text 50. ↵ Bosher JM , Dufourcq P , Sookhareea S , Labouesse M . RNA Interference Can Target Pre-mRNA: Consequences for Gene Expression in a Caenorhabditis elegans Operon . Genetics . 1999 ; 153 : 1245 – 56 . OpenUrl Abstract / FREE Full Text 51. ↵ Guang S , Bochner AF , Burkhart KB , Burton N , Pavelec DM , Kennedy S . Small regulatory RNAs inhibit RNA polymerase II during the elongation phase of transcription . Nature . 2010 ; 465 : 1097 – 101 . OpenUrl CrossRef PubMed Web of Science 52. ↵ Vos JC , van Luenen HG , Plasterk RH . Characterization of the Caenorhabditis elegans Tc1 transposase in vivo and in vitro . Genes Dev . 1993 ; 7 : 1244 – 53 . OpenUrl Abstract / FREE Full Text 53. ↵ Sijen T , Plasterk RHA . Transposon silencing in the Caenorhabditis elegans germ line by natural RNAi . Nature . 2003 ; 426 : 310 – 4 . OpenUrl CrossRef PubMed Web of Science 54. ↵ Fischer SEJ . Activity and Silencing of Transposable Elements in C. elegans . DNA . 2024 ; 4 : 129 – 40 . OpenUrl CrossRef 55. ↵ Wan G , Yan J , Fei Y , Pagano DJ , Kennedy S . A conserved NRDE-2/MTR-4 complex mediates nuclear RNAi in Caenorhabditis elegans . Genetics . 2020 ; 216 : 1071 – 85 . OpenUrl Abstract / FREE Full Text 56. ↵ Billi AC , Fischer SEJ , Kim JK . Endogenous RNAi pathways in C. elegans . WormBook . 2014 ; 1 – 49 . 57. ↵ McCloskey A , Taniguchi I , Shinmyozu K , Ohno M . hnRNP C tetramer measures RNA length to classify RNA polymerase II transcripts for export . Science . 2012 ; 335 : 1643 – 6 . OpenUrl Abstract / FREE Full Text 58. ↵ Liu N , Dai Q , Zheng G , He C , Parisien M , Pan T . N(6)-methyladenosine-dependent RNA structural switches regulate RNA-protein interactions . Nature . 2015 ; 518 : 560 – 4 . OpenUrl CrossRef PubMed 59. ↵ Laski FA , Rio DC , Rubin GM . Tissue specificity of Drosophila P element transposition is regulated at the level of mRNA splicing . Cell . 1986 ; 44 : 7 – 19 . OpenUrl CrossRef PubMed Web of Science 60. ↵ Ghanim GE , Rio DC , Teixeira FK . Mechanism and regulation of P element transposition . Open Biol . 2020 ; 10 : 200244 . OpenUrl CrossRef PubMed 61. ↵ Tabara H , Sarkissian M , Kelly WG , Fleenor J , Grishok A , Timmons L , et al. The rde-1 gene , RNA interference, and transposon silencing in C. elegans. Cell . 1999 ; 99 : 123 – 32 . OpenUrl CrossRef 62. ↵ Brenner S . The genetics of Caenorhabditis elegans . Genetics . 1974 ; 77 : 71 – 94 . OpenUrl Abstract / FREE Full Text 63. ↵ Arribere JA , Bell RT , Fu BXH , Artiles KL , Hartman PS , Fire AZ . Efficient marker-free recovery of custom genetic modifications with CRISPR/Cas9 in Caenorhabditis elegans . Genetics . 2014 ; 198 : 837 – 46 . OpenUrl Abstract / FREE Full Text 64. ↵ Ghanta KS , Mello CC . Melting dsDNA donor molecules greatly improves precision genome editing in Caenorhabditis elegans . Genetics . 2020 ; 216 : 643 – 50 . OpenUrl Abstract / FREE Full Text 65. ↵ Concordet J-P , Haeussler M . CRISPOR: intuitive guide selection for CRISPR/Cas9 genome editing experiments and screens . Nucleic Acids Res . 2018 ; 46 : W242 – 5 . OpenUrl CrossRef PubMed 66. ↵ Schwartz ML , Jorgensen EM . SapTrap, a Toolkit for High-Throughput CRISPR/Cas9 Gene Modification in Caenorhabditis elegans . Genetics . 2016 ; 202 : 1277 – 88 . OpenUrl Abstract / FREE Full Text 67. ↵ Dickinson DJ , Pani AM , Heppert JK , Higgins CD , Goldstein B . Streamlined Genome Engineering with a Self-Excising Drug Selection Cassette . Genetics . 2015 ; 200 : 1035 – 49 . OpenUrl Abstract / FREE Full Text 68. ↵ Frøkjær-Jensen C , Davis MW , Ailion M , Jorgensen EM . Improved Mos1-mediated transgenesis in C. elegans . Nat Methods . 2012 ; 9 : 117 – 8 . OpenUrl CrossRef PubMed Web of Science 69. ↵ Frøkjær-Jensen C , Davis MW , Sarov M , Taylor J , Flibotte S , LaBella M , et al. Random and targeted transgene insertion in Caenorhabditis elegans using a modified Mos1 transposon . Nat Methods . 2014 ; 11 : 529 – 34 . OpenUrl CrossRef PubMed 70. ↵ Schindelin J , Arganda-Carreras I , Frise E , Kaynig V , Longair M , Pietzsch T , et al. Fiji: an open-source platform for biological-image analysis . Nat Methods . 2012 ; 9 : 676 – 82 . OpenUrl CrossRef PubMed Web of Science 71. ↵ Eide D , Anderson P . The gene structures of spontaneous mutations affecting a Caenorhabditis elegans myosin heavy chain gene . Genetics . 1985 ; 109 : 67 – 79 . OpenUrl Abstract / FREE Full Text 72. ↵ Livak KJ , Schmittgen TD . Analysis of Relative Gene Expression Data Using Real-Time Quantitative PCR and the 2−ΔΔCT Method [Internet] . Methods . 2001 . p. 402 – 8 . Available from : doi: 10.1006/meth.2001.1262 OpenUrl CrossRef PubMed Web of Science 73. ↵ Waterhouse AM , Procter JB , Martin DMA , Clamp M , Barton GJ. Jalview Version 2--a multiple sequence alignment editor and analysis workbench . Bioinformatics . 2009 ; 25 : 1189 – 91 . OpenUrl CrossRef PubMed Web of Science 74. Troshin PV , Procter JB , Barton GJ . Java bioinformatics analysis web services for multiple sequence alignment--JABAWS:MSA . Bioinformatics . 2011 ; 27 : 2001 – 2 . OpenUrl CrossRef PubMed Web of Science 75. ↵ Edgar RC . MUSCLE: multiple sequence alignment with high accuracy and high throughput . Nucleic Acids Res . 2004 ; 32 : 1792 – 7 . OpenUrl CrossRef PubMed Web of Science 76. ↵ UniProt Consortium . UniProt: The universal protein knowledgebase in 2025 . Nucleic Acids Res . 2025 ; 53 : D609 – 17 . OpenUrl CrossRef PubMed View the discussion thread. Back to top Previous Next Posted October 10, 2025. Download PDF Supplementary Material Data/Code Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. 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