Function, Targets, and Evolution of Caenorhabditis elegans piRNAs

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Science  03 Aug 2012:
Vol. 337, Issue 6094, pp. 574-578
DOI: 10.1126/science.1220952

Secondary Endogenous Small and Interfering

In many eukaryotes, Piwi proteins bind small noncoding Piwi-interacting RNAs (piRNAs) that function to silence transposons in the germ line and protect the germ line from transposable element–driven recombination and mutation. Bagijn et al. (p. 574, published online 14 June; see the Perspective by Xiol and Pillai) show that in the nematode, Caenorhabditis elegans, a messenger RNA (mRNA) that contains a piRNA target sequence gives rise to a second, downstream class of small RNAs known as secondary endogenous small interfering RNAs, or endo siRNAs. These endo siRNAs map to the vicinity of the piRNA complementary sequence in the mRNA target and depend on both Piwi and on factors involved in the related RNA interference pathway for their genesis, but not on the Piwi slicer activity. Mapping the endo siRNAs reveals that piRNAs can target imperfectly matched targets and that piRNAs target a subset of both transposons and endogenous genes for silencing.


Piwi-interacting RNAs (piRNAs) are small RNAs required to maintain germline integrity and fertility, but their mechanism of action is poorly understood. Here we demonstrate that Caenorhabditis elegans piRNAs silence transcripts in trans through imperfectly complementary sites. Target silencing is independent of Piwi endonuclease activity or “slicing.” Instead, piRNAs initiate a localized secondary endogenous small interfering RNA (endo-siRNA) response. Endogenous protein-coding gene and transposon transcripts exhibit Piwi-dependent endo-siRNAs at sites complementary to piRNAs and are derepressed in Piwi mutants. Genomic loci of piRNA biogenesis are depleted of protein-coding genes and tend to overlap the start and end of transposons in sense and antisense, respectively. Our data suggest that nematode piRNA clusters are evolving to generate piRNAs against active mobile elements. Thus, piRNAs provide heritable, sequence-specific triggers for RNA interference in C. elegans.

The Piwi-piRNA pathway has an evolutionarily conserved role in germline transposon silencing in animals. Caenorhabditis elegans encodes two Piwi family proteins, PRG-1 and PRG-2, although PRG-2 has likely little or no function (1, 2). PRG-1 and piRNA expression is restricted to the male and female germ line. The piRNAs of C. elegans are 21 nucleotides in length with a 5′ uracil (21U-RNAs) (14). In C. elegans piRNAs have a sequence motif, situated ~40 base pairs (bp) upstream of each piRNA locus, that is thought to be required for piRNA biogenesis (2, 3). A challenge in the field is to understand the mechanism(s) by which piRNAs act on their targets. Proposed functions for Piwi-piRNA complexes include the RNA interference (RNAi)–like slicing of RNA transcripts (57), transcript deadenylation (8), and de novo DNA methylation (9, 10). Here we identify the targeting mechanism and the endogenous targets of C. elegans piRNAs.

We generated C. elegans strains carrying green fluorescent protein (GFP)–histone H2B fusion transgenes into which we inserted a short sequence complementary to that of an endogenous piRNA (21UR-1) or its reverse complement, hereafter referred to as the piRNA sensor and control sensor, respectively (see fig. S1 and supplementary text) (11). Whereas the control sensor expressed nuclear, chromatin-associated GFP throughout germline development, GFP expression was silenced in animals carrying the piRNA sensor transgene (Fig. 1, A and B). piRNA sensor silencing was dependent on prg-1. In contrast, prg-2 did not show an effect (fig. S2 and supplementary text). An independent sensor for the unrelated endogenous piRNA 21UR-1349 confirmed these results (Fig. 1, C and D).

Fig. 1

A single antisense piRNA site is sufficient for target silencing in vivo. (A) Fluorescence microscopy (GFP-H2B) and differential interference contrast (DIC) images of adult hermaphrodites. Scale bar, 20 μm. (B) Flow cytometry analysis of control sensor strain (green) and piRNA sensor strain in wild-type (red) or prg-1 (n4357) mutant background (blue) as in (A). (C) Germline GFP-H2B expression of the 21UR-1349 piRNA sensor. (D) Flow cytometry analysis of the 21UR-1349 piRNA sensor strain in wild-type (red) and prg-1 mutant (blue). (E) Profiles of small RNA high-throughput sequencing reads with unique match to the sensor relative to the target site (indicated in gray). Colors correspond to 5′ nucleotides as indicated in the color key in (F). Positive and negative y axes correspond to antisense and sense reads, respectively. (F) Length and 5′ nucleotide identity of small RNAs antisense to the piRNA sensor in wild type. (G) Northern blot of total RNA. Probes were against piRNA 21UR-1, a piRNA sensor-specific 22G-RNA, and the PRG-1–independent endo-siRNA siR26-263. (H) Quantitative reverse transcription (qRT)–PCR of primary piRNA sensor transcript and mRNA. Data were normalized to wild-type transcript levels. Error bars are SEMs.

We analyzed small RNA populations in the sensor strains by high-throughput sequencing (Fig. 1E) (11). We detected a set of small RNAs that map unambiguously to the piRNA sensor mRNA in antisense orientation, mostly within ~20 bp of the piRNA target site. These small RNAs were predominantly 22 nucleotides in length with a 5′ guanosine, characteristic features of secondary endo-siRNAs, also referred to as 22G-RNAs (Fig. 1F). 22G-RNAs represent the most abundant class of endogenous small RNAs in C. elegans, are RNA-dependent RNA polymerase products, and have a 5′ triphosphate (1214). The set of 22G-RNAs that map in close proximity to the piRNA target site was dependent on prg-1 and absent in the control sensor (Fig. 1E). We validated the high-throughput sequencing data using Northern blotting for an abundant 22G-RNA mapping to the piRNA sensor (Fig. 1G). As endo-siRNA silencing can involve posttranscriptional regulation of mRNAs (15) or cotranscriptional gene regulation (16), we tested whether the piRNA sensor was regulated at either level. We find that both primary transcript and mRNA are up-regulated in prg-1 mutants, consistent with cotranscriptional or a combination of post- and cotranscriptional regulation of the piRNA (Fig. 1H).

To investigate the role of endo-siRNAs in piRNA-mediated gene silencing, we screened a set of 24 siRNA pathway genes. Mutants were crossed into the piRNA sensor and assayed for transgene GFP expression through fluorescent microscopy (Fig. 2A and fig. S3, A and B), flow cytometry (fig. S3, C and D), Northern blotting (fig. S4), or high-throughput sequencing (fig. S5). We found that a specific subset of siRNA pathway genes is required for piRNA-mediated silencing and encodes pathway components, including three putative helicases (MUT-7, DRH-3, MUT-14), two RNA-dependent RNA polymerases (EGO-1 and RRF-1), the enzymes that generate secondary siRNAs, and several “worm”-specific Argonaute proteins (WAGOs) (17) (fig. S6, A and B). Although several strains with mutations in multiple WAGOs are defective in piRNA sensor silencing, hrde-1/wago-9 is the only single-mutant WAGO strain defective in piRNA sensor silencing (Fig. 2A and fig. S3). PRG-1 is necessary for both piRNA expression and sensor-specific 22G-RNA expression. In contrast, the identified siRNA pathway genes act downstream of piRNA expression and are required only for the expression of piRNA sensor-derived 22G-RNAs (figs. S4 and S5). We conclude that a specific endo-siRNA pathway acts as a downstream effector of the piRNA pathway.

Fig. 2

A specific endo-siRNA pathway acts downstream of and is required for piRNA-mediated silencing. (A) piRNA sensor expression in siRNA pathway mutants as in Fig. 1A. (B) A second 21UR-1 piRNA sensor strain (cherrysensor) expressing mCherry-H2B in prg-1 (n4357).

The Ping-Pong piRNA amplification loop in insects (18) and vertebrates requires endonuclease or slicing activity for biogenesis of at least a subset of piRNAs (7, 19). PRG-1 contains an evolutionarily conserved DDH motif (catalytic triad) that confers endonuclease or slicing activity to some Argonaute superfamily proteins (20), and recombinant PRG-1 appears to have some slicing activity in vitro (figs. S7 and S8 and supplementary text). To test the requirement for PRG-1 catalytic activity in vivo, we generated two transgenes expressing wild-type GFP–PRG-1 or GFP–PRG-1 DAH mutant fusion proteins (fig. S9). In prg-1 mutants, both wild-type and DAH mutant GFP–PRG-1 were sufficient to rescue piRNA expression (fig. S4). To address whether the catalytic triad of PRG-1 is required for piRNA-mediated silencing of targets, we generated a second 21UR-1 piRNA sensor strain (cherrysensor) expressing mCherry-H2B. Although the cherrysensor was desilenced in a prg-1 mutant background, both wild-type GFP–PRG-1 and GFP–PRG-1 DAH restored silencing of the cherrysensor to the same extent (Fig. 2B). This was also true in the prg-1;prg-2 double mutants (fig. S10). In addition, both wild-type and DAH mutant GFP–PRG-1 rescued the fertility defects of prg-1 mutants (fig. S11). Finally, we generated a panel of mutated piRNA sensors and find that two mismatches are tolerated throughout the target sequence for PRG-1–dependent sensor silencing, including mismatches at positions 10 and 11 that are required for slicing (fig. S12). We conclude that PRG-1 slicing is not required in vivo.

To investigate whether piRNAs target endogenous transcripts, we considered piRNA matches in the C. elegans genome, allowing for up to three mismatches. For 16,003 piRNAs, we identified a total of 681,746 sites (additional data tables S1 and S2). We found that PRG-1–dependent 22G-RNAs localize in close proximity to imperfect piRNA matches, recapitulating our observations for the piRNA sensor (Fig. 3A and fig. S13). To assess how many of these represent functional target sites, we compared genomic matches of piRNAs to those of matched control sequences. In wild-type animals, approximately 4.2, 2.6, 2.0, and 1.7% of the 0, 1, 2, and 3 mismatch sites exhibit unambiguously mapping 22G-RNAs, corresponding to an enrichment of 1.6, 1.4, 1.5, and 1.2 compared to control sites, respectively (Fig. 3B). In prg-1 mutants, the percentage of genomic piRNA matches with 22G-RNAs was comparable to that of control sites. The distribution of piRNA matches (and those with 22G-RNAs) across the genome resembled that of controls, except for an enrichment of perfect piRNA matches in the two piRNA clusters on chromosome IV (figs. S14 and S15).

Fig. 3

piRNAs initiate a localized secondary siRNA response against endogenous transcripts. (A) Average profiles of collapsed small RNAs mapping uniquely to imperfect genomic piRNA matches (one to three mismatches) in wild-type (left) and prg-1 mutant (right). Top and bottom panels to the right of each profile illustrate characteristics of antisense and sense small RNAs, respectively. (B) Number of genomic piRNA matches (left) and percentage of matches with uniquely mapping 22G-RNAs in wild type (middle) and prg-1 (right). Black and white bars correspond to piRNAs and matched controls, respectively. Bars for control sequences indicate medians, error bars the range of values obtained for 20 cohorts of control sequences. Numbers above bars indicate the fold-difference between piRNAs and controls. (C) Difference in 22G-RNAs mapping uniquely within 20 bp of genomic piRNA matches between prg-1 and wild type. Shown are boxplots of the difference in 22G-RNA reads after square root transformation (box indicates interquartile range, plot extends from 5th to 95th percentile). Asterisks indicate statistical significance (P < 0.001, two-sided Wilcoxon rank-sum test). (D) As in (C) with genomic piRNA matches grouped according to motif score of complementary piRNA (as proxy for abundance).

To further investigate mismatch tolerance of piRNA targeting, we considered genomic piRNA matches with up to five mismatches. We observed that levels of PRG-1–dependent 22G-RNAs decreased with an increasing number of mismatches (Fig. 3C). Levels of PRG-1–dependent 22G-RNAs were greater at sites with four compared to five mismatches, suggesting that in some cases, sites with up to four mismatches can be sufficient for the synthesis of 22G-RNAs. We also observed that levels of 22G-RNAs depend on piRNA abundance (Fig. 3D).

We identified candidate endogenous targets by searching for antisense piRNA matches in annotated protein-coding genes, pseudogenes, and transposons, allowing for up to three mismatches (additional data tables S3 to S5). We found that in prg-1 mutant animals, 22G-RNAs antisense to candidate targets showed a stronger reduction at target sites (within 20 nucleotides) compared to regions distant from target sites (fig. S16). Microarray analysis showed that mRNA expression differences between prg-1; prg-2 mutant and wild-type animals were more strongly correlated with changes in 22G-RNAs at target sites compared to the whole transcript (fig. S16).

We ranked transposons and protein-coding genes by the decrease in target-site–associated 22G-RNA density in prg-1 mutant compared to wild-type animals and examined individual candidate targets (Fig. 4, A and B). Transposase mRNA from Tc3 is desilenced in prg-1 mutants, and Tc3 exhibits PRG-1–dependent 22G-RNAs against its terminal inverted repeats (TIRs) but no matching piRNAs (1, 2). We identified three piRNAs with imperfect complementarity to the consensus sequence of Tc3, all of which map to the TIRs, suggesting that these piRNAs target Tc3 elements in trans. We chose five transposable elements (CEREP1A, MARINCE1, TURMOIL1, Chapaev-2_CE, and LINE2H_CE) and six protein-coding genes (bath-45, zfp-1, C18H2.2, nfm-1, Y75B8A.19, and pan-1) with a strong reduction in target-site–associated 22G-RNAs for analysis by quantitative reverse transcription–polymerase chain reaction (qRT-PCR) (Fig. 4, C to E). Six of these candidates showed statistically significant increased expression in prg-1 mutants (P < 0.05, two-tailed t test). We tested the requirement of an intact catalytic triad and found that both wild-type and DAH mutant GFP–PRG-1 restored silencing of zfp-1 (F54F2.2b) in prg-1 mutants, whereas an independent isoform lacking target-site–associated 22G-RNAs (F54F2.2a) showed no change in expression (Fig. 4E). In addition, expression of a PRG-1–dependent 22G-RNA against zfp-1 (F54F2.2b) was restored by both wild-type and DAH mutant GFP–PRG-1 (fig. S4). Thus, piRNAs target endogenous transcripts in trans for silencing, independently of slicing activity.

Fig. 4

Endogenous piRNA targets and piRNA evolution. (A) Candidate transposon targets ranked by change in 22G-RNA density at target sites between prg-1 and wild type. Transposons selected for qRT-PCR validation are in red. Antisense 22G-RNA profiles are shown for selected elements with target sites indicated above each profile as explained in the color key. (B) Candidate protein-coding targets as in (A). (C) qRT-PCR analysis of candidate transposon targets with fold-changes normalized to actin. Error bars are SEMs, asterisks denote P < 0.05 (two-sided t test). (D) qRT-PCR analysis of candidate protein-coding targets. (E) qRT-PCR analysis of targeted (F54F2.2b) and nontargeted (F54F2.2a) transcripts from the zfp-1 locus. Data were normalized to wild type. (F) Enrichment and depletion of genomic piRNA matches overlapping features of interest. Red and blue indicate increased or reduced number of matches for piRNAs compared to control sequences, respectively. Asterisks indicate statistical significance (adjusted empirical P < 0.05). Start and end refer to the first and last 50 bp of the annotated feature, respectively. Pseudogene annotation was only available for C. elegans. (G) piRNA matches against start (left) and end (right) of DNA transposons. Profiles indicate the number of transposon subfamilies with a perfect piRNA match in at least one full-length genomic copy. Positive (blue) and negative (red) y axes correspond to sense and antisense matches, respectively. Dashed lines correspond to maximal allowed distance between an upstream sequence motif and piRNA 3′ end.

Our findings raise the question of how piRNAs and target sites arise during evolution. The majority of the 16,003 C. elegans piRNAs (96%) map to unique locations in the genome and are depleted of protein-coding genes (2). We confirmed that piRNA loci are depleted of protein-coding genes in both C. elegans and the related nematode species C. briggsae when compared to loci of matched control sequences (Fig. 4F). However, we did not observe a depletion of pseudogenes or transposons. piRNA loci showed a trend for depletion at the start and end of full-length DNA transposons in antisense and sense, respectively, and an inverse trend for enrichment at the start and end in sense and antisense, respectively (Fig. 4F). This signature suggests recent DNA transposon integrations downstream of instances of the sequence motif thought to be required for piRNA biogenesis. Such integrations may result in the birth of a piRNA either sense to the 5′ end or antisense to the 3′ end of the transposon. In the latter case, the new piRNA has the potential to target and silence the mobile element. Indeed, the observed distances between piRNA locus and start or end of the transposon are consistent with transposon insertions downstream of existing sequence motifs (Fig. 4G and fig. S17). A similar signature was observed in C. briggsae (Fig. 4G and fig. S18) even though neither piRNAs nor transposable elements are conserved between the two species. When considering imperfect piRNA matches, we observed a depletion antisense to protein-coding genes (Fig. 4F and fig. S19), suggesting that mRNA targeting is often detrimental and piRNAs and sites with potential for mRNA silencing undergo negative selection.

Our data demonstrate that C. elegans piRNAs silence endogenous transcripts by triggering a secondary siRNA response (fig. S20). Although we find that individual mRNAs are piRNA targets, the physiological roles of piRNA-mediated gene regulation remain to be explored. RNA interference pathways can silence repetitive elements regardless of their sequence content but rely on the formation of double-stranded RNA. The piRNA pathway may provide an alternative defense mechanism that is heritable and sequence-specific based on the evolution of new piRNAs against active mobile elements. Secondary siRNA amplification ensures effective silencing of abundant targets, resembling the Ping-Pong piRNA amplification cycle in other species.

Supplementary Materials

Materials and Methods

Supplementary Text

Figs. S1 to S21

Tables S1 to S3

References (2130)

Additional Data Tables S1 to S5

References and Notes

  1. Materials and methods are available as supplementary materials on Science Online.
  2. Acknowledgments: We are grateful to J. Ahringer and E. Jorgensen for providing reagents. A.S. was supported by the Human Frontier Science Program and the Herchel-Smith Fund. N.J.L. was supported by a Ph.D. studentship from the Wellcome Trust (UK). E.-M.W. was supported by a Ph.D. studentship from the Herchel-Smith Fund. S.B. and M.J.S. were funded by the Canadian Institutes of Health Research (CIHR). M.J.S. is a New Investigator from CIHR. This work was supported by Cancer Research UK and the Wellcome Trust. Small RNA sequence data and Affymetrix gene expression data were submitted to the Gene Expression Omnibus under accession no. GSE37433. Detailed methods can be found in the supplementary materials.
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