Introduction
Since August Weismann (1834–1914) formulated the distinction between innate and acquired
characteristics at the end of the 19th century, the debate relating to the inheritance
of acquired traits has raised many controversies in the scientific community (Weismann,
1891; Bateson, 1919; Haig, 2006). August Weismann himself theoretically rejected this
type of hereditability, arguing that, even though environmental stimuli could provoke
adaptive responses in the somatic lineage, no evidence suggested that these changes
could be communicated to the germline (Weismann, 1891). However, a number of epigenetic
phenomena involving RNA, histone modification, or DNA methylation in many organisms
have renewed interest in this area (Varmuza, 2003; Haig, 2006; Daxinger and Whitelaw,
2012). Paramutation is a prime example. In this phenomenon, a silenced allele can
act in trans on a homologous sequence to cause stable and heritable silencing. This
newly silenced allele can now itself act in a paramutagenic fashion to silence other
alleles. Paramutation has been described in multiple species, and it seems likely
that small RNAs play a key role in the process, although the full mechanisms involved
still remain unclear (Stam and Mittelsten Scheid, 2005; Chandler, 2010; Suter and
Martin, 2010).
C. elegans has emerged as a key model for the analysis of several related pathways
that regulate genes via small RNAs. C. elegans is well suited to the analysis of multigenerational
effects, due to its short generation time (∼3 days) and the ease with which they can
be maintained under tightly controlled experimental conditions. In eukaryotes, 20–30
nucleotide (nt) RNAs bound to Argonaute (AGO) protein cofactors are the effectors
of a number of gene regulation pathways (Carmell et al., 2002). The discovery of the
process of RNA interference (RNAi) has been a major milestone (Fire et al., 1998).
While 21–22 nt small interfering RNAs (siRNAs) are the small RNA effectors of RNAi,
RNAi can be induced by injection of long double-stranded RNA (dsRNA) or by providing
dsRNA environmentally in the food of C. elegans (Timmons et al., 2001). In both instances,
dsRNA is processed by the RNase Dicer to give rise to primary siRNAs. RNAi effects
are generally systemic (soma and germline) and are observed in the F1 generation,
but the latter requires the generation of secondary siRNAs (Grishok et al., 2000;
Pak and Fire, 2007; Sijen et al., 2007; Gu et al., 2009). Secondary siRNAs represent
the most abundant class of endogenous small RNAs in C. elegans, are RNA-dependent
RNA polymerase products, have a 5′ triphosphate, and are predominantly 22 nt in length
with a 5′ guanosine (22G-RNAs). Secondary siRNA pathways and RNA-dependent RNA polymerases
(RdRPs) have not been found in vertebrates or Drosophila, but have been found in many
other organisms, including nematodes, plants, fungi, and viruses. Secondary siRNA
pathways in C. elegans are complex, can involve many different AGO proteins, and are
only partly understood (Yigit et al., 2006).
Several studies have reported inheritance of environmental RNAi beyond the F1 generation
(Fire et al., 1998; Grishok et al., 2000; Vastenhouw et al., 2006; Alcazar et al.,
2008; Gu et al., 2012). In one transgenerational paradigm, small RNA inheritance and
histone H3K9me3 marks were observed for at least two generations (Gu et al., 2012).
In addition, transgenerational inheritance of viral immunity (Rechavi et al., 2011)
and longevity (Greer et al., 2011) were recently reported for C. elegans. These data
suggest that the biological roles of transgenerational inheritance could be diverse
but remain largely speculative. In addition, whether this transmission involves transgenerationally
transmitted RNAs or modifications of chromatin is still controversial.
Piwi-interacting RNAs (piRNAs) are distinct from siRNAs and have an evolutionarily
conserved role in transposon silencing in the germline in many animals, including
nematodes (Malone and Hannon, 2009; Bagijn et al., 2012). C. elegans encodes two Piwi
clade, AGO superfamily proteins, PRG-1 and PRG-2, although PRG-2 has likely little
or no function (Batista et al., 2008; Das et al., 2008; Bagijn et al., 2012). C. elegans
piRNAs are absent in prg-1 mutant animals, which exhibit fertility defects. PRG-1
and piRNA expression is restricted to the male and female germline (Batista et al.,
2008; Das et al., 2008; Bagijn et al., 2012). The piRNAs of C. elegans are 21 nucleotides
in length with a 5′ uracil (21U-RNAs) (Ruby et al., 2006; Batista et al., 2008; Das
et al., 2008; Wang and Reinke, 2008). C. elegans piRNAs derive from two broad clusters
on chromosome IV (Ruby et al., 2006) and act in trans to regulate endogenous targets
in the germline (Bagijn et al., 2012).
Here, we report how transgenerational inheritance of environmental RNAi and the piRNA
pathway converge on the same germline nuclear RNAi/chromatin pathway. Both nuclear
RNAi factors and chromatin regulators are essential for silencing. This pathway can
elicit a long-term epigenetic memory for more than 24 generations. Once established,
the initial silencing trigger is no longer required.
Results
A Reporter-Based System to Investigate Transgenerational Gene Silencing in C. elegans
To genetically dissect multigenerational gene silencing in C. elegans, we developed
a heritable environmental RNAi paradigm. Taking advantage of the recent advance in
technologies to generate single-copy intrachromosomal transgenes in C. elegans (Frøkjaer-Jensen
et al., 2008, 2012), we generated a reporter transgene expressing a histone-GFP fusion
protein in the germline of C. elegans (Figure 1A and Figure S1 available online).
The use of such a defined artificial locus combines the ability to map small RNA populations,
which was previously not possible using multicopy transgenes (Vastenhouw et al., 2006),
with a high-throughput, quantifiable approach that was not possible using an endogenous
locus (Alcazar et al., 2008). Eliciting environmental RNAi by feeding transgenic animals
with bacteria expressing dsRNA corresponding to the GFP mRNA results in gene-specific
silencing of this GFP transgene (P0, Figures 1A and 1B), as expected (Timmons et al.,
2001). Transfer of these animals to a neutral environment results in a high level
of silenced animals in the F1 generation, again as expected (Figure 1B) (Fire et al.,
1998; Grishok et al., 2000). Furthermore, silencing of the transgene is maintained
for at least four additional generations in a subpopulation of animals. We quantified
this phenomenon in thousands of animals for each generation using flow cytometry and
found that inheritance of transgene silencing was maintained in more than 60% of animals
for at least four generations (Figures 1C and S2). We conclude that we have established
a reporter-based paradigm for the investigation of transgenerational inheritance,
the “heritable RNAi defective,” or “Hrde,” sensor.
Multigenerational Gene Silencing Is Associated with Continued Small RNA Expression
As the mechanisms of transgenerational inheritance are currently not understood, we
first asked whether the Hrde sensor silencing that we observed is due to posttranscriptional
regulation of mRNA or (co-)transcriptional gene regulation. Using quantitative RT-PCR,
we tested whether Hrde sensor silencing in the F2 generation affected either. We were
able to robustly detect both primary transcript (pre-mRNA) and Hrde transgene mRNA.
However, mRNA levels were significantly repressed (p < 0.05) in silenced animals as
compared to nonsilenced animals (Figure 1D). pre-mRNA levels showed a similar trend.
These data suggest that posttranscriptional mechanisms of silencing are required in
the Hrde paradigm. We postulated that Hrde sensor transcript availability might result
in continued small RNA pathway activity in silenced animals. Therefore, we profiled
small RNAs using high-throughput sequencing from animals undergoing environmental
RNAi (P0), control RNAi (P0), or at the F4 generation after RNAi. Small RNA libraries
were prepared using protocols that did not necessitate the presence of a 5′ monophosphate
to capture primary and secondary siRNAs. We detected abundant sense and antisense
small RNAs during environmental RNAi (P0 generation) (Figure 1E). These small RNAs
had a peak length of 21–22 nt and little bias for the 5′-most nucleotide and likely
represent Dicer cleavage products (primary siRNAs) (Figure 1F). In contrast, four
generations later, only antisense small RNAs remain with the characteristic signature
of secondary siRNAs (22 nucleotide length with a 5′ guanosine bias, 22G RNAs). Given
that each generation represents at least a hundred-fold dilution in volume (with more
than 200 offspring generated by each hermaphrodite), these secondary siRNAs must be
generated de novo in each generation. Animals undergoing control RNAi displayed a
peak of small RNAs homologous to cloning sequences flanking the GFP minigene. These
are homologous to cloning sequences that are present in the RNAi vectors and have
no apparent effect on Hrde sensor activity (Figure 1C).
Multigenerational Gene Silencing and piRNA Silencing Depend on Common Nuclear Factors
We recently reported that piRNA-mediated silencing in the C. elegans germline results
in secondary siRNA-dependent silencing of a “piRNA sensor” and endogenous piRNA targets
(Bagijn et al., 2012). Thus, piRNA-mediated silencing might converge on a common downstream
multigenerational gene silencing pathway. To this end, we carried out forward genetic
screens to identify genes required for either phenomenon using the Hrde and piRNA
sensors. Using these two distinct sensors (Figure S1), one silenced by a single endogenous
piRNA (piRNA sensor) and the other silenced by heritable environmental RNAi (Hrde
sensor), we identified, mapped, and cloned new alleles of three known genes in small
RNA pathways: nrde-2, nrde-4, and hrde-1/wago-9 (Table 1). Next, combining forward
genetic screens with a candidate gene approach, we were surprised to identify a total
of eight small RNA or chromatin pathway genes to be required (Table 1). For example,
the Hrde sensor was desilenced in nrde-2, hrde-1/wago-9, and set-25 mutants (Figure 2A
and Table 1). The products of all of these genes are either known to be, or are predicted
to be, nuclear. NRDE-2 is a conserved protein involved in nuclear RNAi that is expressed
in the nucleus (Guang et al., 2010); SET-25 is a putative histone H3 lysine-9 methyltransferase
with a C-terminal SET domain. To our knowledge, this is the first time that a histone-modifying
enzyme has been identified as required for multigenerational inheritance. HRDE-1/WAGO-9
is an Argonaute protein. Using immunostaining, we show that it is expressed in the
germline (Figure 2B), where it localizes to the nucleus (Figures 2C and 2D). NRDE-2
was recently shown to be important in a similar inheritance paradigm (Gu et al., 2012).
However, some genes that were previously reported to be involved in transgenerational
effects appeared not to be required for our transgenerational inheritance paradigm,
including hda-4, mrg-1 (Vastenhouw et al., 2006) or spr-5, lsd-1, and amx-1 (Katz
et al., 2009) (Table 1). For the piRNA sensor, aside from proteins that were defined
in Hrde screens such as NRDE-2, HRDE-1/WAGO-9, and SET-25, we also identified additional
nuclear small RNA components and chromatin factors, including NRDE-1, NRDE-4, SET-32,
and one of the C. elegans heterochromatin protein 1 (HP1) orthologs, HPL-2 (Table
1 and Figure S3). We conclude that there exists a common and specific nuclear RNAi/chromatin
pathway in the germline that is required for environmentally induced heritable RNAi-
and piRNA-induced silencing.
The Nuclear RNAi/Chromatin Silencing Pathway Acts Downstream of Small RNA Expression
in Gene Silencing
To establish a hierarchy in the silencing pathways described here, we asked whether
nuclear RNAi/chromatin components are upstream or downstream of secondary siRNA expression
and/or stability. First, we analyzed small RNA expression in the Hrde sensor paradigm
in wild-type and a nrde-2 mutant background. In both cases, we find abundant 22G secondary
siRNAs that map to the Hrde sensor (Figure 3A). Thus, NRDE-2 is not required for secondary
siRNA generation. Next, we tested a chromatin factor using the piRNA sensor. The HP-1
ortholog HPL-2, but not HPL-1, is required for silencing of the piRNA sensor (Table
1 and Figure 3B). Therefore, we asked whether secondary siRNAs are expressed and stable
in hpl-2 mutant animals. Using northern blotting, we show that the piRNA 21UR-1 and
a piRNA-sensor-specific 22G RNA (siR22G-1) are dependent on the Piwi protein PRG-1
(Figure 3C). However, both RNAs are present in hpl-2 and hpl-2; hpl-1 mutant backgrounds,
although possibly at reduced levels for siR22G-1. These observations are in agreement
with similar observations made for siRNAs in S. pombe lacking Swi6/HP1 (Bühler et al.,
2006). In addition, we analyzed endogenous targets of the piRNA pathway that we recently
identified (Bagijn et al., 2012). We generated small RNA libraries from wild-type
and prg-1 or hpl-2 mutant animals. 22G secondary siRNAs at endogenous piRNA targets
bath-45 and zfp-1 are dependent on PRG-1, but not HPL-2 (Figures 3D and 3E). Again,
we observed some reduction in RNA levels, consistent with a positive interaction between
nuclear RNAi and chromatin regulation. We conclude that the nuclear RNAi/chromatin
pathway described here is not essential for secondary siRNA expression or stability.
Multigenerational Gene Silencing and piRNA Silencing Does Not Spread into the Soma
As the nuclear RNAi/chromatin pathway that we describe here utilizes small RNAs, it
might act in trans on transcripts that share significant sequence similarity. Indeed,
using the piRNA sensor, we were able to test this directly. The piRNA sensor is under
the transcriptional control of a germline-specific promoter (mex-5). Silencing of
the piRNA sensor is established through an endogenous piRNA (21-UR-1) with perfect
complementarity to a corresponding sequence in the piRNA sensor (Bagijn et al., 2012).
A cross of the piRNA sensor strain to a different transgenic strain with a ubiquitously
expressed GFP transgene that is not regulated by piRNAs (Figure 4A) results in dominant
silencing of both transgenes in the germline of heterozygous F1 animals (Figure 4B),
likely via a process termed transitive RNAi (Alder et al., 2003). Thus, the nuclear
RNAi/chromatin pathway can silence in trans. We postulate that this effect is mediated
via secondary siRNAs. As exogenous and endogenous RNAi are systemic in C. elegans
(Fire et al., 1998; Winston et al., 2002), we therefore wondered whether the germline
nuclear RNAi/chromatin silencing pathway that we describe here could transcend the
germline/soma boundary. We do not find this to be the case, as GFP expression in the
trans-heterozygous animals (dpy-30::his-58::gfp::tbb-2/piRNA sensor) described above
remains unaffected in the soma (Figure 4B). We made the same observation using the
Hrde-1 sensor and another somatic transgene in an analogous experiment (data not shown).
We conclude that, though the nuclear RNAi/chromatin pathway that we describe here
can be vertically transmitted, it does not trigger systemic RNAi. This is consistent
with recent work demonstrating that secondary siRNAs are not systemically transmitted
in the soma of C. elegans (Jose et al., 2011). We note that results based on multicopy
transgenes that possibly involve dsRNA intermediates could be different from those
reported here (Jose et al., 2011).
piRNAs Can Trigger Long-Term Multigenerational Gene Silencing
Our data demonstrate that environmentally induced multigenerational gene silencing
and piRNA silencing converge on a common germline silencing pathway. Can a piRNA therefore
trigger multigenerational gene silencing? To address this question, we carried out
genetic crosses in which we removed PRG-1 and thereby piRNA function from the piRNA
sensor strain (Das et al., 2008; Bagijn et al., 2012). In these circumstances, the
piRNA trigger is removed but silencing might be maintained. In a cross of animals
homozygous for the piRNA sensor (GFP silenced) with an animal homozygous for the piRNA
sensor but in a prg-1 mutant background (GFP expressed), we generated F1 animals homozygous
for the piRNA sensor but heterozygous for the recessive mutation in prg-1 (Figure S4).
Such animals are GFP silenced for several generations, as expected. These heterozygous
animals segregate progeny that are homozygous, heterozygous, or wild-type with respect
to prg-1. We observed piRNA sensor reactivation in prg-1 homozygous mutants or their
immediate offspring. Because all piRNAs are eliminated in prg-1 mutants (Batista et al.,
2008; Das et al., 2008), these data suggested that a piRNA trigger may be required
to maintain multigenerational silencing memory.
Next, we recreated a piRNA sensor strain that was mutant for prg-1 by outcrossing
the piRNA sensor and then performing several crosses using mutations that cause visible
phenotypes to mark the positions of prg-1 or the piRNA sensor transgene (see Experimental
Procedures). Unexpectedly, 11 prg-1; piRNA sensor strains failed to reactivate the
piRNA sensor (n = 8 or 3 independent strains created per trial for 2 trials) (Figures
5A and 5B). GFP expression of these prg-1; piRNA sensor strains failed to materialize
even though many successive generations were scored, which were last analyzed at F16,
F17 (three strains), F20, and F24 (six strains) generations. We also observed that
silencing can become PRG-1 independent using a second piRNA sensor construct integrated
on a different chromosome (the piRNA mCherry sensor; Figure S5). We conclude that
germline silencing can persist for many generations even in the absence of a piRNA
trigger. It is of interest to note that all crosses that led to trigger-independent
maintenance of silencing involved the piRNA sensor transgene being heterozygous for
3–5 generations due to outcrossing.
In contrast to multigenerational silencing of piRNA sensor transgenes in the absence
of prg-1, mutation of nrde-1 (yp4 or yp5) or mutation of nrde-2 (gg95) triggered reactivation
of outcrossed piRNA sensors (n = 3, 2, and 3, respectively, independently isolated
F3 or F4 strains scored) (Figure 5C). All nrde-2 mutant lines expressed bright GFP
from F3 onward. Of five piRNA sensor; nrde-1 lines, three lines expressed weak GFP
signal in all germ cells in the F3 generation, whereas germ cells of all animals scored
in piRNA sensor; nrde-1 lines were uniformly positive for a weak GFP signal by the
F4 generation. We conclude that nuclear small RNA factors are required to maintain
the silenced state over many generations, whereas the piRNA trigger that initiates
silencing becomes dispensable if the silent locus is outcrossed multiple times.
A Tudor domain protein RSD-6 and a novel protein RSD-2 have previously been shown
to be required for RNAi responses to environmental dsRNA triggers that target genes
expressed in the germline and are proficient for RNAi to some somatic targets, possibly
due to dose-dependent RNAi defects (Tijsterman et al., 2004; Merritt et al., 2008;
Zhang et al., 2012). The strong germline RNAi defects of rsd-6 or rsd-2 suggested
that they could function to promote systemic spreading of RNAi from soma to germline
(Tijsterman et al., 2004). To determine where rsd-6 functions to promote germline
RNAi, single-copy rsd-6 transgenes driven by the germline-specific pgl-3 promoter
or by the ubiquitous promoter dpy-30 were created (Frøkjaer-Jensen et al., 2008; Han
et al., 2008). Both transgenes rescued an rsd-6 mutant for the response to dsRNAs
targeting the germline-expressed genes pop-1 or par-6 (Figure S6), indicating that
RSD-6 functions in a cell-autonomous manner within the germline. We created rsd-6;
piRNA sensor and piRNA sensor; rsd-2 strains using outcrossed sensor transgenes and
observed that these strains were GFP negative when initially created and for many
generations thereafter (Figure 5C). These results suggest that piRNA sensor silencing
may not depend on systemic RNAi effects (possibly mediated by expression of dsRNA
in somatic cells). They also suggest that the response to dsRNA generated in germ
cells is unlikely to promote sensor silencing (Tabara et al., 1999; 2002).
Discussion
Here, we show that piRNA and environmental RNAi pathways converge on a common germline
nuclear RNAi/chromatin pathway. This pathway can induce stable, multigenerational
inheritance. Previous work found evidence for inheritance of small RNAs, chromatin,
or both in related transgenerational inheritance paradigms (Burton et al., 2011; Rechavi
et al., 2011; Gu et al., 2012). However, here we demonstrate that both small RNA and
chromatin factors are essential for multigenerational inheritance and do not act redundantly
(Table 1 and Figures 2, 3, S3). We also show that small RNA biogenesis occurs upstream
of nuclear RNAi and chromatin factors (Figures 3). Recent work has proposed that somatic
nuclear RNAi acts at the level of transcriptional elongation (Guang et al., 2010).
These observations opened the possibility that chromatin changes observed in transgenerational
inheritance paradigms (Gu et al., 2012) might simply be correlative without being
functional in silencing. However, our data show that chromatin factors, such as HPL-2
and SET-25/32, are required (Table 1 and Figures 3 and S3). SET-25/32 are putative
histone H3K9me3 methyltransferases. This histone modification, a hallmark of silenced
chromatin, has been correlated with small RNA-mediated transgene silencing (Shirayama
et al., 2012 [this issue of Cell]; Gu et al., 2012) and is enriched on the Hrde sensor
reported here (data not shown). In addition, multigenerational silencing of transgenes
is promoted by HPL-2 and SET domain proteins (Shirayama et al., 2012). Though this
related study did not examine the requirement for SET-25 or SET-32, it did find that
MES-4, a histone H3K36 methyltransferase that participates in silencing of the X chromosome
(Bender et al., 2006; Rechtsteiner et al., 2010), is also required for multigenerational
inheritance. These observations suggest that the chromatin states involved in multigenerational
silencing might be complex and could include a hierarchy, which merits further investigation.
We have summarized a model of our current understanding of this pathway in Figure 6.
Multicopy versus Single-Copy Transgenes
Multicopy transgenes, intra- or extrachromosomal, are generally efficiently silenced
in the germline of C. elegans (Kelly et al., 1997). This has been interpreted as an
example of the RNAi machinery distinguishing self from nonself (Vastenhouw and Plasterk,
2004). In this model, repetitive DNA such as endogenous transposable elements or multicopy
transgenes would give rise to dsRNA that is processed by Dicer to generate siRNA triggers
to induce silencing. As the pathways silencing multicopy transgenes and transposable
elements share common factors, this phenomenon is of biological interest. However,
it has also been a major technical roadblock for researchers studying germ cell biology
who rely on reproducible transgene expression in the germline. The advent of MosSCI
technology to produce single-copy transgenes has the promise to overcome this problem
(Frøkjaer-Jensen et al., 2008, 2012). Interestingly, we and others observed that,
in some cases, individual transgenes remain silenced even when present as single,
intrachromosomal entities (N.J.L. and E.A.M., unpublished data) (Frøkjaer-Jensen et al.,
2008). Indeed, an accompanying paper reports a collection of MosSCI transgenes that
remain silenced (Shirayama et al., 2012). Generating MosSCI transgenes in animals
in which germline nuclear RNAi pathways are impaired, such as mut-7, or “curing” silenced
transgenes by outcrossing first to germline RNAi mutants strains and then back to
wild-type often results in loss of transgene silencing (N.J.L., A.S., and E.A.M.,
unpublished data). These results suggest that the original RNAi model of multicopy
transgene silencing needs to be revised. Indeed, there appears to be no requirement
for dsRNA intermediates in the silencing phenomena reported here either, as factors
required for dsRNA-induced RNAi in the germline such as RSD-2 and RSD-6 (Figure 5
and Table 1) or RDE-1 and RDE-4 (Shirayama et al., 2012) are dispensable for single-copy
transgene silencing.
Self versus Nonself
How does C. elegans detect single-copy transgenes and target them for silencing in
the germline, or how does the animal distinguish self from nonself? The answer might
lie in a combination of three factors: scanning germline gene expression by the piRNA
pathway (nonself RNA recognition), licensing of germline transcripts (self RNA recognition),
and unpaired genomic DNA in meiosis. Based on this and related work, we propose that
the piRNA pathway can detect transgenes as sources of foreign RNA (nonself) and initiates
targeted silencing of these transgenes (Bagijn et al., 2012; Lee et al., 2012 [this
issue of Cell]). The piRNA pathway is perfectly suited for this task, as it provides
a diverse and large set (∼15,000) of small RNA triggers that are mismatch tolerant
but do not depend on dsRNA generation (Ruby et al., 2006; Batista et al., 2008; Das
et al., 2008; Wang and Reinke, 2008; Bagijn et al., 2012). Furthermore, endogenous
germline transcripts are generally depleted in piRNA target sites (Bagijn et al.,
2012). In addition, germline licensing pathways might act in the opposite manner to
protect bona fide germline transcripts. A recent study reported such a phenomenon
in mutants of the fem-1 locus in C. elegans (Johnson and Spence, 2011). In this case,
maternal transcripts were required to overcome silencing of an endogenous locus in
a mutant background. Furthermore, the germline Argonaute CSR-1 associates with secondary
siRNAs that map to many germline-expressed genes without inducing silencing (Claycomb
et al., 2009). Interestingly, CSR-1-bound 22G RNAs appear to match abundantly to single-copy
transgenes that evade silencing (Shirayama et al., 2012). Taken together, a balance
of nonself recognition by the PRG-1/piRNA pathway and self recognition by licensing
factors such as the CSR-1 pathway might determine the outcome of gene expression in
the germline. This model helps to explain the apparent discrepancy between the facultative
multigenerational inheritance that we observe here in our piRNA sensor and the obligatory
multigenerational inheritance observed for a related piRNA sensor in a parallel study
(Lee et al., 2012; Shirayama et al., 2012). Differences in the composition of the
sensors, e.g., the inclusion of the coding region of the his-58 gene in our sensor,
might tip the PRG-1/CSR-1 pathway balance. However, this model fails to explain the
ability of our piRNA sensor to be silenced or active depending on its multigenerational
ancestry. In our crosses, the piRNA sensor became stably silenced when present in
a heterozygous state for several generations (Figures 5 and S5). We propose that unpaired
chromatin that has been subjected to silencing by a piRNA trigger can be subjected
to an additional layer of silencing during meiosis that then makes the original piRNA
trigger dispensable (Figure 6). Unpaired DNA silencing responses have been observed
in C. elegans in the case of the unpaired X chromosome (Kelly et al., 2002; Bean et al.,
2004) and for a mutant fem-1 locus (Johnson and Spence, 2011) and has also been found
in other organisms (Hynes and Todd, 2003; Lee, 2005; Matzke and Birchler, 2005). Establishment
of heritable silent chromosome domains that can be robustly maintained in the absence
of the original piRNA trigger could be relevant to populations in which sources of
piRNAs are polymorphic and may evolve rapidly in response to novel transposons or
retroviruses.
Related Phenomena in Other Phyla
The core molecular pathway described here is reminiscent of related (co-) transcriptional
pathways both in yeasts and plants (Moazed, 2009). Many yeasts and all plants and
nematodes share key factors, such as the RNA-dependent RNA polymerases involved in
secondary siRNA generation. Though transgenerational phenomena have been reported
in many animals, including humans (Hitchins et al., 2007), this class of polymerases
and secondary siRNAs appears to be absent in Drosophila and vertebrates. However,
it is interesting to note that Drosophila and vertebrates have a more complex piRNA
system that includes an amplification loop termed “ping-pong,” which could function
in a manner analogous to secondary siRNA pathways (Brennecke et al., 2007; Gunawardane
et al., 2007). Despite differences in details of piRNA and secondary siRNA systems,
common downstream silencing mechanisms may exist.
One, Few, or Many Generations?
Transgenerational phenomena have been observed over one or multiple generations (Grishok
et al., 2000; Vastenhouw et al., 2006; Alcazar et al., 2008; Burton et al., 2011;
Rechavi et al., 2011; Gu et al., 2012). In some cases, inheritance is stochastic;
in others, Mendelian. Here, we report that piRNAs can trigger silencing that lasts
for more than 20 generations (Figures 5A, 5B, and S5). Although maintenance of this
memory is observed in 100% of offspring, establishment of strong piRNA-independent
memory is not obligatory (Figure S5) and only occurs if a silent locus is heterozygous
for several generations. This is reminiscent of ubiquitous yet stochastic inactivation
of repetitive germline transgenes in many organisms, including C. elegans. Our study
of transgenes targeted by an endogenous piRNA may recapitulate the fate of transposons
that are transmitted in rare horizontal transfer events, in which a single transposon
insertion could be subjected to dual layers of silencing, as the locus would likely
remain heterozygous for a number of generations before potentially becoming fixed.
It will be of great interest to identify the factor(s) that determines these distinct
states of silencing.
Experimental Procedures
Genetics
C. elegans were grown under standard conditions at 20°C unless otherwise indicated.
The food source used was E. coli strain HB101 (Caenorhabditis Genetics Center, University
of Minnesota, Twin Cities, MN, USA). Bleaching followed by starvation-induced L1 arrest
was used to generate synchronized cultures. The wild-type strain was var. Bristol
N2 (Brenner, 1974). All strains used are listed in Table S1. For details about genetic
crosses, see Supplemental Information.
Transgenics
To generate transgenic animals, germline transformation was performed as described
(Mello and Fire, 1995). Injection mixes contained 2–20 ng/μl of MosSCI plasmid and
5–10 ng/μl of marker plasmid DNA (see Supplemental Information for details). Single-copy
transgenes were generated by transposase-mediated integration (MosSCI), as described
(Frøkjaer-Jensen et al., 2008, 2012).
COPAS Biosort Analysis
A COPAS Biosort instrument (Union Biometrica, Holliston, MA, USA) was used to simultaneously
measure length (time of flight), absorbance (extinction), and fluorescence. Data handling
and analysis were performed using FlowJo (Tree Star, Inc.) and R.
Extended Experimental Procedures
Genetic Crosses
Testing nrde- or prg-1-Mediated Suppression of mjIs144
Marker double mutants rol-6; dpy-17 or unc-13; rol-6 were first created to make prg-1;
mjIs144 or mjIs144, nrde-2 or mjIs144; nrde-1 strains. To facilitate making strains
with mjIs144, the unc-119(ed3) mutation was first removed from SX1316 by crossing
rol-6 e187 / + males with single SX1316 hermaphrodites, selecting for F1 with Rol
F2, singling many non-Rol siblings, selecting for F2 with Rol F3 but no Unc F3, and
then selecting against rol-6 to obtain mjIs144 homozygotes. unc-13; mjIs144 and mjIs144;
dpy-17 strains were then created by crossing rol-6; dpy-17 or unc-13; rol-6 with N2
wild-type males, crossing F1 rol-6; dpy-17 / +; + or unc-13; rol-6 / +; + males with
single mjIs144 hermaphrodites, selecting for F1 that segregated both marker mutations,
and then selecting for F2 that lacked rol-6 and then unc-13 or dpy-17 F3. Heterozygous
prg-1 / unc-13 males stock were maintained by backcrossing heterozygous males with
unc-13 homozygotes. Heterozygous nrde-1 / dpy-17 males stock were maintained by backcrossing
heterozygous males with dpy-17 homozygotes. prg-1; rol-6 strains were crossed with
prg-1 / unc-13 males, and F1 males were crossed with unc-13; mjIs144 hermaphrodites,
and F1 with both marker mutations were transferred for several generations to obtain
strains that lacked Unc or Rol mutations, which were prg-1; mjIs144 strains. rol-6;
nrde-1 animals were crossed with nrde-1 / dpy-17 males, F1 males were crossed with
mjIs144; dpy-17 hermaphrodites, and F1 with both marker mutations were transferred
for several generations to obtain strains that lacked Rol or Dpy mutations, which
were mjIs144; nrde-1. nrde-2 is tightly linked to the piRNA sensor (mjIs144) on chromosome
II, so dpy-10, piRNA sensor, unc-4 strains were first created, and then F2 Dpy-non-Unc
recombinants from dpy-10, mjIs144, unc-4 / +, +, nrde-2 F1 were singled, allowed to
self, and genotyped for the presence of the piRNA sensor. Three of 6 F2 Dpy-non-Unc
recombinants possessed the piRNA sensor, and F4 progeny of F3 homozygotes for piRNA
sensor, nrde-2 displayed uniformly GFP. To construct piRNA sensor; rsd-2 strains,
rsd-2; rol-6 and unc-24; mjIs144 strains were created using a rol-6; unc-24 strain.
rol-6; rsd-2 hermaphrodites were then crossed with rsd-2 males. Male offspring were
crossed with unc-24; mjIs144 hermaphrodites. F1 from this cross were singled. F2 with
an unaffected phenotype but with both rol-6 and unc-24 siblings were singled. F2 that
segregated neither unc-24, nor rol-6 offspring were selected, producing strains homozygous
for both rsd-2 the piRNA sensor. To construct rsd-6; piRNA sensor strains, rsd-6;
rol-6 and dpy-5 unc −55; mjIs144 strains were created using a dpy-5 unc-55; rol-6
strain. rsd-6; dpy-10 unc-4 hermaphrodites were then crossed with rsd-6 males. Male
offspring were selected and crossed with dpy-5 unc-55; mjIs144 hermaphrodites. F1
were singled and those which produced dpy-5 unc-55 and dpy-10 unc-4 F2 were selected.
Unaffected F2 were singled from these plates. Finally, F2 which segregated neither
dpy-5 unc-55 nor dpy-10 unc-4 were selected. Independent strains that were homozygous
for both rsd-6 and the piRNA sensor were tested for GFP fluorescence.
Genotyping
SNP Mapping
SNP mapping of mutations was performed as described previously (Bruinsma et al., 2008).
Primer sequences are available on request.
Other Genotyping
In order to verify that the prg-1; mjIs144 strains had both the prg-1 n4357 allele
and the mjIs144 transgene, DNA was extracted from a recently starved plate of each
prg-1; mjIs144 strain. PCR was performed using Bam-GFP-FW and Bam-GFP 2 Stop-RV primers
to confirm the presence of the transgene. DNA from wild-type worms and mjIs144 worms
were used as controls. The PCR products revealed an approximately 800 bp band indicating
the presence of the GFP DNA sequence in the mjIs114 control as well as all in eight
prg-1; mjIs144 strains, but not in the wild-type control. PCR using n4357 FW and n4357
RV primers was used to verify the presence of the prg-1 n4357 allele. DNA from wild-type
worms and prg-1 n4357 worms were used as controls. The PCR products revealed a long
band non mutant band for wild-type but only a shorter band for the n4357 deletion
control as well as for all eight prg-1; mjIs144 strains, confirming that the prg-1;
mjIs144 n4357 strains were homozygous for n4357.
Transgenics
rsd-6 Rescue Transgenes
Rescue constructs were prepared using MultiSite Gateway Three-Fragment Vector Construction
Kit (Invitrogen). A PCR product corresponding to the coding region of rsd-6 was amplified
from the fosmid WRM064bA10 (Source Bioscience) and subcloned into pDONR 221. Ubiquitously
expressed dpy-30 promoter and the 3′UTR of rsd-6 were amplified from N2 wild-type
genomic DNA and subcloned into pDONR P4-P1R and pDONR P2R-P3 vectors, respectively.
Germline-specific pgl-3 promoter is a gift from Kyle Wang. Fragments for promoter,
gene and 3′UTR were combined into pCFJ150 vector (Frøkjaer-Jensen et al., 2008) including
Caenorhabditis briggsae unc-119(+) gene. All pDONR subclones were sequenced to confirm
lack of mutations, and att recombination sites were sequenced in the final promoter:gene:3′UTR
constructs prior to microinjection. Primers were following; attB1-RSD6FW, ggggacaagtttgtacaaaaaagcaggctatgaatgaaaaagagctggcggatt;
attB2-RSD6RV, ggggaccactttgtacaagaaagctgggttcagataaagacgtctttgatattc; attB4-dpy30p-FW,
ggggacaactttgtatagaaaagttggtctattctcacacctctcc; attB1-dpy30p-RV, ggggactgcttttttgtacaaacttgcttggtttttgctcgatttct;
attB2-rsd6-3U-FW, ggggacagctttcttgtacaaagtggacttcaaatcatgtttctatctaaa; attB3-rsd6-3U-RV,
ggggacaactttgtataataaagttgtctcatgtatattgtttgatgtgaa.
Transgenes were injected into rsd-6(pk3300) I, ttTi5605 II (Mos1); unc-119(ed3)III
and the integrants were obtained using the ‘Mos1 excision-induced transgene-instructed
gene conversion’ method (Frøkjaer-Jensen et al., 2008). Single copy inserts at the
ttTi5605 locus were fully sequenced to confirm a lack of deletions or other mutations
prior to use of transgenes for complementation tests. RNAi feeding for rsd-6 mutants
with or without transgenes was performed by placing L4 larvae on P1 RNAi plates, transferring
to P2 RNAi plates at 24 hr, removing adults from P2 plates at 48 hr, and scoring plates
for unhatched embryos 20 hr after adults were removed.
Microscopy
Differential interference contrast (DIC) and fluorescence imaging was performed using
standard methods using an AxioImager A1 upright microscope (Zeiss, Jena, Germany).
Images were captured using an ORCA-ER digital camera (Hamamatsu, Hamamatsu, Japan)
and processed using OpenLabs 4.0 software (Improvision, Coventry, UK). For Figure 5
GFP expression was monitored by mounting adult worms on agar pads and monitoring 10
to 20 worms per strain using a Nikon E800 epifluorescence microscope and a 40X Plan
Fluor objective.
qRT-PCR
RNAi inheritance assay was performed as described above. Silenced animals were sorted
using the biosorter and used to generate the F2 generation. Silenced F2 worms were
sorted and collected into Trizol ®. Control worms were grown alongside under identical
conditions. RNA extraction was performed using standard protocols. cDNA was synthesized
from 5 μg total RNA using Superscript III reverse transcriptase (Invitrogen) with
random hexamers. qRT-PCR was performed using a ABI7300 Real Time PCR system (Applied
Biosystems). Primers sequences are as follows: nascent transcript F: TCTGTCAGTGGAGAGGGTGA;
nascent transcript R: TTTAAACTTACCCATGGAACAGG; mRNA F: CTACCTGTTCCATGGCCAAC; mRNA
R: GGCATGGCACTCTTGAAAAA.
RNAi Inheritance Assay
RNAi bacteria was inoculated into LB Broth containing Ampicillin (50 μg/ml) for 6 hr
at 37°C with shaking. Bacterial cultures were then seeded on to NGM plates containing
IPTG (1 mM) and Carbenicillin (25 μg/ml) and grown overnight at room temperature.
Adult animals were placed on the bacteria and the cultures grown at 20°C for 4 days.
GFP fluorescence in the germline and oocytes of adult offspring was assayed using
either a large particle biosorter (Union Biometrica) or a fluorescent microscope (Kramer
scientific). Ten (biosorter) or one (microscope) silenced adults were selected and
placed on HB101 growth plates to produce the next generation. This procedure was repeated
for up to five generations.
Hrde EMS Screen
After EMS treatment following standard protocols, synchronized P2 larvae (L1) were
placed onto GFP RNAi plates for 3 days at 20°C. Adult RNAi P0 animals (EMS P2s) were
individually picked onto standard HB101 NGM plates and grown at 20°C for 4 days. The
resultant F1 (EMS P3) animals were screened for GFP phenotype and any plates with
a substantial number of GFP expressing animals kept for secondary screening. Secondary
screening was performed as described above for the RNAi inheritance assay. Screening
was performed in 8 batches with 600 P2s individualized each time.
piRNA Sensor EMS Screen
F2 or F3 offspring of mutagenized worms were sorted using a Copas Biosort Large-Particle
Sorter as described in (Bagijn et al.). Worms sorted as ‘GFP ON’ were re-selected
for sensor de-silencing using an FBS10 Fluorescence Microscope System (Kramer Scientific).
Preparation of Genomic DNA for Whole-Genome Sequencing
We used a pellet of approximately 60-100 μl of packed mixed stage worms and froze
at −80°C. For extraction of genomic DNA we followed the Quiagen DNeasy Blood and Tissue
kit (animal tissue spin protocol) including an RNase treatment step. We eluted DNA
in 2x 200 μl buffer and quantified using the QuBit dsDNA BR assay kit (Invitrogen).
Prior to library preparation we cleaned 150 ng of genomic DNA using the DNA Clean&
Concentrator-5 Kit (Zymo Research Corp.). We eluted in 15 μl pre-heated water and
re-measured DNA concentration using the QuBit dsDNA HS DNA assay kit.
Whole-Genome Library Preparation and Sequencing
We prepared libraries from genomic DNA using the EPICENTRE Biotechnologies Nextera
DNA Sample Prep Kit. Tagmentation was performed in HMW Buffer and PCR-amplified libraries
were purified either using the DNA Clean & Concentrator-5 Kit (Zymo Research Corp.)
or Agencourt AMPure XP beads (Beckman Coulter). Library concentration was measured
using the QuBit dsDNA HS DNA assay kit, then fragment length and distribution were
determined with the Agilent High Sensitivity DNA assay (Agilent Technologies Inc.).
Whole-genome sequencing of simplex libraries was performed on either the Genome Analyzer
IIx or the High-Seq 2000 Illumina platform. Sequencing data were analyzed as described
previously (Bagijn et al.).
Small RNA Library Generation
P0 and F4 Samples
Total RNA was isolated using the mirVana miRNA isolation kit (Ambion). The total RNA
samples were separated on denaturing 15% polyacrylamide gels and 18-30 nt length species
selected. Following poly-A tailing, small RNAs were treated with tobacco acid pyrophosphatase
(Epicenter). Adapters were then ligated to the 5′ phosphate of the RNA. First-strand
cDNA synthesis was performed using an oligo(dT)-adaptor primer and the M-MLV reverse
transcriptase. The resulting cDNAs were PCR-amplified to about 20-30 ng/μl in 18-20
PCR cycles using a high fidelity DNA polymerase. cDNA were purified using the Macherey
& Nagel NucleoSpin Extract II kit. Libraries were sequenced at CRI using an Illumina
GA2x instrument.
P0 and F4 Samples
Total RNA was isolated using TRIsure reagent (Bioline). Total RNA was treated with
RNA 5′ polyphosphatase (Epicenter). cDNA libraries were then prepared following the
TruSeq Small RNA protocol (Illumina). Libraries were sequenced on a MiSeq machine
(Illumina).
Computational Analysis of Small RNA High-Throughput Sequencing Data
Data processing and analyses were performed using custom Perl scripts and the statistical
programming environment R. Fastq sequence reads with missing bases or barcodes not
matching any of the expected sequences were excluded (if applicable). Reads were trimmed
by removing 5′ barcodes, 3′ adapters or 3′ As depending on the protocol used for library
generation. Inserts with length 18-30 nucleotides were collapsed to unique sequences,
retaining the number of reads for each sequence. The C. elegans genome (assembly WS190/ce6)
and refGene annotation were downloaded from the UCSC Genome Browser website (http://genome.ucsc.edu/)
(Kent et al., 2002; Fujita et al., 2011). Perfect matches to the reference genome
and transgene constructs (if applicable) were identified using bowtie in -v alignment
mode, allowing for multiple matches (Langmead et al., 2009).
Antibody Generation and Immunostaining
The anti-HRDE-1/WAGO-9 polyclonal antisera were generated using Genomic Antibody Technology
(GAT) by SDIX (Newark, USA). In brief, DNA immunization was done using cDNA encoding
amino acids 12 to 161 of HRDE-1/WAGO-9. Polyclonal sera were subsequently affinity-purified
and tested for specificity in Western blotting (data not shown) and immunostaining.
Immunostainings were performed on dissected gonads. Animals were dissected in 10 mM
Tetramisol on microscopy slides. After freeze-cracking, the tissue was fixed in ice-cold
Methanol for 20 min and rehydrated by three washes in 1xPBS/0.2% Tween 20 (10 min
each, room temperature).
Primary antibodies were added over night at 4°C in 1xPBS/0.2% Tween 20/1% BSA. Slides
were washed three times in 1x PBS/0.2% Tween 20 (10 min each, room temperature). Secondary
antibodies were added for one hour at 37°C. The secondary antibody mix contained 4′,6-diamidino-2-phenylindole
(DAPI) at 100 ng/ml final concentration. Slides were washed three times in 1xPBS/0.2%
Tween 20 (10 min each, room temperature) and mounted using Vectashield. We carried
out differential interference contrast (DIC) and confocal fluorescence imaging using
standard methods and using an Olympus FluoView FV1000 upright microscope using 40x
and 63x objective magnification. Images were processed using the FluoView image software
(Olympus) and ImageJ (version 1.43u). Antibodies and concentrations used are: rabbit
anti-HRDE-1/WAGO-9 1:4000, mouse OIC1D4 (Developmental Studies Hybridoma Bank, University
of Iowa) 1:100, mouse anti-α-Tubulin (clone DM1A, SIGMA) 1:10,000, Alexa Fluor 488
anti-rabbit IgG, 1:1000, Alexa Fluor 568 anti-mouse IgG, 1:1000 (both Life Technologies).