Yamanaka et al. Mobile DNA 2014, 5:22
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REVIEW

Open AccesspiRNA clusters and open chromatin structure
Soichiro Yamanaka1, Mikiko C Siomi2 and Haruhiko Siomi1*
Abstract
Transposable elements (TEs) are major structural components of eukaryotic genomes; however, mobilization of TEs
generally has negative effects on the host genome. To counteract this threat, host cells have evolved genetic andepigenetic mechanisms that keep TEs silenced. One such mechanism involves the Piwi-piRNA complex, which represses
TEs in animal gonads either by cleaving TE transcripts in the cytoplasm or by directing specific chromatin modifications
at TE loci in the nucleus. Most Piwi-interacting RNAs (piRNAs) are derived from genomic piRNA clusters. There has beenremarkable progress in our understanding of the mechanisms underlying piRNA biogenesis. However, little is known
about how a specific locus in the genome is converted into a piRNA-producing site. In this review, we will discuss apossible link between chromatin boundaries and piRNA cluster formation.
Keywords: Transposable elements, Piwi, piRNA, piRNA cluster, Chromatin boundary

Review
Background

Large fractions of eukaryotic genomes comprise transposable elements (TEs), which are repetitive DNA elements that can mobilize to take up new chromosomal
locations within a genome. TEs act as insertional mutagens that can alter gene expression or rearrange chromosomes. Therefore, they can cause disease and may
even drive evolution [1-4]. TEs are diverse in sequenceand in the way they transpose [5,6]. They possess a limited gene set of their own, but use the gene expression
machinery of their host to thrive in the genome. DNA
transposons move by a “cut-and-paste” mechanism, inwhich they are excised from one genomic site andinserted into a new location using their own transposase.
Therefore, generally, the copy number of DNA transposons in a genome does not expand. By contrast, retrotransposons use a “copy-and-paste” mechanism to propagate
their copies through RNA intermediates. Retrotransposons are transcribed from the genome, reverse transcribedand integrated into a new location, in a process mediatedby a transposon-encoded reverse transcriptase. Retrotransposons are distinguished by their DNA sequence topology and mechanism of transposition: those that possesslong terminal repeats (LTRs), such as gypsy, and those that
* Correspondence: awa403@z2.keio.jp
1
Department of Molecular Biology, Keio University School of Medicine,
35 Shinanomachi Shinjuku-ku, Tokyo 160-8582, Japan
Full list of author information is available at the end of the article

do not (non-LTRs), such as long interspersed repetitive elements (LINEs) and short interspersed repetitive elements(SINEs). Both DNA transposons and retrotransposons
have non-autonomous subtypes and defective copies,
which require the reverse transcriptase and endonuclease supplied by the autonomous type to jump aroundthe genome.
As an example, Drosophila harbors around 100 different TEs, and the only conserved and universal property
shared by them is the ability of transposition [7]. Thus,
the requirements for host cells for repression of TEs areat least two-fold: 1) a mechanism that recognizes such adiverse set of TE types, and 2) a mechanism that distinguishes them from other cellular genes and selectively
targets them for silencing. Recent studies have postulated that host cells have evolved an elaborate silencing
mechanism to meet these two requirements. Host cells
may have taken advantage of the only universal propertyof TEs, their transposition ability to trap them in specificgenomic locations and subject them to a silencing program, which employs small RNA-based immunity to selectively silence homologous elements [8-10]. In animal
gonads, small non-coding RNAs (ncRNAs), termed Piwiinteracting RNAs (piRNAs), mediate TE silencing to ensure genome integrity during germ cell development[11,12]. Most piRNAs are derived from particular genomic
sites termed piRNA clusters, which contain a large number and various types of TEs. Thus, the sequences of piRNAs derived from these clusters are homologous not only

© 2014 Yamanaka et al.; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of theCreative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use,
distribution, and reproduction in any medium, provided the original work is properly credited. The Creative Commons Public
Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in thisarticle, unless otherwise stated.

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to TEs in the clusters, but also to related TEs located elsewhere in the genome and can therefore act as guide molecules to repress TEs in trans. Thus, piRNA clusters are
genetic elements that regulate the activity of TEs. However, relatively little is known about how piRNA clusters
are formed. In this review, we emphasize the role of chromatin boundaries in piRNA cluster formation. To thisend, we briefly review our current knowledge of piRNAs
and piRNA clusters. We then discuss a possible link between chromatin boundaries and piRNA clusters, andpropose some models as to how piRNA clusters areformed in chromatin boundaries.
TE silencing mediated by piRNAs

RNA interference (RNAi) and related pathways are cellular pathways in which small ncRNAs of 20 to 35 nucleotides (nt) guide Argonaute-containing effector complexes,
termed RNA-induced silencing complexes (RISCs), to
RNA targets by means of base-pairing, and promote theinactivation of homologous sequences [13-16]. They havebeen shown to suppress the activity of TEs in plants andanimals. In animal germline cells, piRNAs of 24 to 35 nt
are produced and loaded onto germline-specific Argonaute proteins (termed PIWI proteins) to form piRNAinduced silencing complexes (piRISCs). Genetic analysesof Drosophila PIWI genes (ago3, aubergine/aub, and piwi)
have revealed that mutations in these genes affect germline development [17-20]. TEs are deregulated in mutant
ovaries defective in these genes, suggesting a model inwhich TE overexpression and mobilization triggers DNA
damage signaling-dependent defects in an early step in thegerm cell patterning cascade [21].
Unlike other small silencing RNAs such as microRNAs(miRNAs) and small interfering RNAs (siRNAs), piRNAs
in most animals are processed in a Dicer-independent
manner from single-stranded precursors, which are transcribed mostly from genomic piRNA clusters [22,23]. A
large number of genes have been identified to functionin piRNA biogenesis [24]. In the Drosophila genome,

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142 regions have been identified as piRNA clusters [22].
Although these sites account for less than 5% of the assembled genome, over 90% of all sequenced piRNAs canbe derived from these regions [25]. The piRNA clusters
cover chromosomal regions of several to hundreds of kilobases, and they contain TEs that are mostly inactive
copies or truncated fragments, arranged in a nested
fashion [22]. Among all the piRNA clusters in Drosophila, the flamenco locus produces a major fraction of piRNAs in somatic support cells in the ovary [25]. Thislocus was originally discovered as a regulator of the activity of the gypsy, idefix, and ZAM TEs [26,27]. piRNAs
from this cluster, which spans about 150 kb, are derived
from one DNA strand only, most likely through unidirectional transcription oriented in the anti-sense direction to most TEs in the locus (Figure 1). This provides amolecular basis of why Piwi, the only PIWI protein
expressed in ovarian somatic cells, loads with piRNAs
that are anti-sense-oriented relative to TEs. Mutants offlamenco in which the P-element is inserted in the 5′ region and those lacking flamenco partial genomic sequence lose the ability to regulate TEs [22,26,28,29].
These data indicate that the single long transcripts fromthe flamenco locus are processed into piRNAs. This linear biogenesis of piRNAs from precursor transcripts
has been called the ‘primary piRNA processing pathway’ (Figure 2). piRNA maturation and Piwi-piRNAcomplex (Piwi-piRISC) formation occur in the cytoplasm [30]. Piwi-piRISCs are then imported into the
nucleus where they repress TEs in trans at transcriptional level by directing specific histone modifications
to TE loci [31-34]. This suggests that Piwi-piRISCs recruit the relevant enzymes to modify histones at TE
loci. Because depletion of piwi activity rapidly results inderepression of TEs, the TE silencing state requires thecontinual activities of Piwi-piRISCs [30,35,36]. Therefore, Piwi-piRISCs are genetic elements that mediate
and maintain epigenetic chromatin modifications oftarget TE loci.

Figure 1 flamenco, a major Piwi-interacting RNA (piRNA) cluster in somatic support cells of the Drosophila ovary. The flamenco locus
contains a particular family of transposon (boxes with white arrows; the arrows denotes the direction of each transposon) in its transcription unit.
Almost all transposons are truncated and/or inactivated. The direction of the transposons is exclusively anti-sense with regard to transcription inthis region (gray arrow). This region spans about 150 kb, and is thought to behave as a single transcriptional unit.

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Figure 2 Piwi-interacting RNA (piRNA) biogenesis pathway in the Drosophila ovary. (A) Primary piRNA pathway in somatic support cells(cream region surrounding the central egg). The transposon sequence in piRNA clusters (the majority are unistrand clusters; see Figure 5 below)
in somatic support cells is in an exclusively anti-sense orientation with regard to the direction of transcription. The resultant transcripts aretransported to the cytoplasm, recognized, and processed by several factors, including Zucchini, Armi, and Yb. Finally, they are loaded onto
the PIWI protein. (B) The ping-pong amplification loop in germ cells (light blue region). Transcripts from piRNA clusters (mainly dual-strand
clusters; see Figure 5 below) and active transposons are processed into piRNAs by Aub and Ago3. piRNAs from sense transposon transcript
are preferentially loaded onto Ago3, and those from anti-sense transposon transcript are preferentially loaded onto Aub.

Compared with the situation in somatic support cells,
the piRNA biogenesis in germline cells in the fly ovary ismore complex. In contrast to the unidirectional flamenco piRNA cluster, many piRNA clusters in the Drosophila germline are transcribed from both strands, andboth precursor transcripts are processed into piRNAs[22,25]. Therefore, both sense and anti-sense piRNAs
relative to the TE sequences are produced from the clusters. All three PIWI proteins are expressed in the germline, but Piwi is nuclear, and both Aub and Ago3 arecytoplasmic [22,37,38]. Anti-sense precursor transcripts
from dual-stranded piRNA clusters are processed into
anti-sense piRNAs that are loaded onto Aub and Piwi(“primary piRNA processing pathway”). Piwi-piRISCs
then move into the nucleus where they repress TEs,
probably by a mechanism similar to that observed insomatic support cells. Aub-piRISCs, by contrast, remainin the cytoplasm and cleave both sense precursor transcripts from dual-stranded piRNA clusters and transcripts from active TEs, using the small RNA-directedendonuclease or Slicer activity exhibited by PIWI proteins [37]. This cleavage results in the production ofsense piRNAs, which in turn are loaded onto Ago3. This
process initiates a feed-forward amplification loop ofpiRNA production, the so-called “ping-pong cycle”, inwhich sense and anti-sense transcripts of dual-stranded
piRNA clusters and active TEs are reciprocally cleavedby the Slicer activity of Ago3 and Aub [22,37] (Figure 2).
Both Ago3-piRISCs and Aub-piRISCs act catalytically,

and thus the cycle leads to repeated rounds of piRNA
production by consuming both cluster transcripts andTE transcripts, thereby silencing TEs at posttranscriptional levels in the cytoplasm.
The mouse genome encodes three distinct PIWI proteins: MIWI, MIWI2, and MILI. In contrast to Drosophila PIWI proteins, which are expressed in both male andfemale gonads, the expression of mouse PIWI proteins israther restricted to male gonads [39-41]. Male knock-out(KO) mice for each PIWI gene show defects in spermatogenesis and sterility, but female PIWI KO mice are normal[39-41]. Two distinct piRNA populations are present inmouse testes: the pre-pachytene and pachytene piRNA
pools. Pre-pachytene piRNAs are enriched in TE-derived
sequences (approximately 80% of the total), and associatewith MIWI2 and MILI [39]. Pachytene piRNAs, by contrast, have a higher proportion of unannotated sequences,with diminished contribution from TE-derived sequences(approximately 25%) [42-44]. Pachytene piRNAs enter
MILI and MIWI [42-45] (Figure 3). Similar to the case in
Drosophila, both the primary piRNA processing pathway
and the ping-pong cycle operate in mouse testes. MILI
and MIWI accommodate piRNAs from the primarypiRNA processing pathway, but unlike in Drosophila,
mouse primary piRNAs are predominantly sense-oriented
relative to the TE transcripts [11]. It was initially thoughtthat MILI and MIWI2 form a ping-pong amplificationloop, and that anti-sense piRNAs were loaded onto
MIWI2 to form MIWI2-piRISCs [39,46]. However, recentYamanaka et al. Mobile DNA 2014, 5:22
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Figure 3 The Piwi-interacting RNA (piRNA) biogenesis pathway in mouse testis. The piRNA biogenesis pathway in mouse can be
categorized into three modes. MILI is expressed in both prenatal and adult testis. MIWI2 is expressed in prenatal testis and its expression
decreased after birth and is not detectable in adult testis. MIWI is expressed in adult testis. (A) When MILI and MIWI2 are coexpressed in prenatal
testis, the primary piRNA transcript is processed for loading into MILI. The MILI-piRISC can form homotypic ping-pong amplification loop.
MIWI2-associated piRNAs are processed from anti-sense transcripts with the aid of MILI-piRNA-induced silencing complex (piRISC). Therefore,
the production of MIWI2-associated piRNA depends on mature MILI-piRISC. (B) When only MILI protein is expressed in testis, MILI processsense and anti-sense piRNA precursor transcripts. (C) When MILI and MIWI are coexpressed in adult testis, both Piwi proteins process thesense and anti-sense piRNA precursor transcript.

studies have shown that the Slicer activity of MILI is required for the secondary piRNA production, which amplifies MILI-bound piRNAs through an intra-MILI pingpong loop, and generates all MIWI2-bound secondary
piRNAs [45] (Figure 3). In contrast to the cytoplasmic
localization of MILI and MIWI, MIWI2-piRISCs areimported into the nucleus where they direct specific DNA
methylation of TE loci, thereby establishing TE silencing
at the transcriptional level [39,45,47]. However, the Slicer
activity of both MIWI and MILI is still required to maintain TE silencing in the mouse testis after birth, suggestingthat continuous cleavage of TE transcripts by the Slicer
activity is essential to repress TEs in mouse testes [44,45].
piRNA clusters in diverse organisms

TE insertions in Drosophila are mostly located in heterochromatin and proximal heterochromatin-euchromatinboundary zones [22]. Of 142 piRNA clusters identified
in Drosophila, only 7 are in presumed euchromatic regions, while the rest reside within cytologically defined
pericentromeric and telomeric heterochromatin regions.
Within these heterochromatin regions, the piRNA clusters tend to be located near the boundary region between
heterochromatin and euchromatin. Heterochromatin regions in the Drosophila genome can be found at the pericentromeric and subtelomeric regions, and are megabases
in size [48-50]. Their constituent sequences fall intoroughly three categories: tandemly repeated short sequences (satellite DNAs), moderately repetitive elements(such as TEs), and some single-copy genes [48-50]. In the
Drosophila genome, intact and potentially active TEs prevail across the genome, while fragmented or inactive
copies of TEs are strongly enriched in the transition
zones between heterochromatin and euchromatin nearto the centromere, and constitute piRNA clusters [22,50](Figure 4).
Because most piRNAs are derived from piRNA clusters that genetically control the activity of TEs andlargely comprise various types of defective TEs, a modelin which piRNA clusters act as “TE traps” has been proposed [8,51-53]. This model relies on the transposition
ability of TEs for piRNA clusters to passively acquire
new content by chance transposition. TEs that happen
to jump into piRNA clusters can then become fixed byevolutionary selection, and produce corresponding piRNAs and regulate other homologous elements expressedfrom different genomic positions in germ cells.
As mentioned above, two types of piRNA clusters exist
in the Drosophila gonads: unidirectional clusters anddual-stranded clusters. Most piRNA clusters in somatic
support cells are unidirectional, while the predominantfraction of germline piRNA clusters is dual-stranded[22,25] (Figure 5).
An example of a unidirectional piRNA cluster is theflamenco locus, which is located near the pericentromeric heterochromatin boundary of the X chromosome,and contains a large number of truncated or inactivatedTEs. Most of these TEs belong to the gypsy family andYamanaka et al. Mobile DNA 2014, 5:22
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Figure 4 Most Drosophila Piwi-interacting RNA (piRNA) clusters are found near the boundary zone between euchromatin andheterochromatin. The boundary between euchromatin and heterochromatin of Drosophila is gradual rather than acute. Most Drosophila piRNA
clusters exist in the boundary zone between euchromatin and heterochromatin.

are anti-sense-oriented with regard to the polarity oftranscription. This requires the transcription factor Cubitus interruptus, a segment polarity gene that controls anumber of genes, including Hedgehog genes [22,54].
The molecular mechanism that restricts the directionality of transposition into a unistrand piRNA cluster is notwell understood.
A representative dual-stranded cluster is the 42AB
cluster, which spans around 240 kb, near the pericentromeric heterochromatin boundary. However, the orientation of truncated TEs in this cluster is random rather
than uniform, and piRNAs are produced from bothsense and anti-sense strands.
Although many factors that are required for piRNA
production are shared between these two types of clusters, there are some differences between them. Rhino (avariant of heterochromatin protein 1; HP1), Cutoff (ahomolog of the yeast decapping nuclease and transcription termination factor Rai1), and Deadlock (which acts
as a linker between Rhino and Cutoff ), are all requiredfor piRNA production only in germline cells of the oocyte [22,55-57]. Interestingly, most piRNA clusters in
Drosophila are within cytologically defined heterochromatic regions. A recent chromatin immunoprecipitation(ChIP)-sequencing analysis of H3K9me3, the most established marker for heterochromatic regions, revealed thatthe promoter and its surrounding region of flamenco, aunistrand piRNA cluster, is fairly devoid of this repressive

histone mark, which may explain the active transcriptionof the locus by RNA polymerase II [34]. By contrast, thegermline cell-specific dual-strand piRNA clusters, such as
42AB, are coated with H3K9me3, but are still transcriptionally active [55] (see also below).
In the Bombyx mori tissue cultured cell line BmN4, aportion of piRNAs are derived from TEs [58]. piRNA
clusters in BmN4 cells have been shown to have a highlevel of H3K4me3 mark, which is a hallmark of active
transcription [59], suggesting the open nature of silkworm piRNA clusters.
These findings suggest that piRNA clusters are highly
transcribed units within heterochromatic regions, andraise the question of how this kind of special location in
the genome has been selected for piRNA clusters to produce piRNAs.
In the mouse, over 90% of piRNA reads have been
mapped to roughly 100 genomic regions, ranging from a
few kb to over 100 kb in length. Most mouse clusters
show profound strand asymmetry, with reads arisingfrom only one strand within a cluster (unidirectional
cluster). When piRNAs map to both strands within one
piRNA cluster, the transcription units are arranged in adivergent manner (bidirectional cluster) [42,43] and thepiRNA-producing region on one strand does not overlap
with that on the other strand. In prenatal mouse testes,
piRNAs are produced from both strands in the same region(dual-strand cluster) [39] (Figure 5). Recent comprehensive

Figure 5 Three types of Piwi-interacting RNA (piRNA) cluster. (A) Unistrand piRNA cluster; piRNAs are produced from only one genomic
DNA strand. (B) Dual-strand piRNA cluster; piRNAs are produced from both strands of the same genomic region. (C) Bidirectional piRNA cluster;
two unistrand piRNA clusters are arranged in a divergent manner.

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deep sequencing analysis of postnatal mouse testes reveals
that the transcription factor A-MYB drives pachytene
piRNA production, suggesting a model in which a specifictranscription factor engages in transcription of most piRNA
clusters [60,61]. It should be noted that A-MYB is not specific for piRNA clusters, but rather has a number of target
genes, suggesting that A-MYB has been co-opted to drive
transcription of piRNA clusters. This also raises the questionof what might be the difference between the A-MYB binding sites that direct piRNA production and the A-MYB
binding sites that produce mRNAs but not piRNAs. piRNA
clusters have been identified in other mammals including
primates [62]. Synteny analysis has revealed conservation in
the genomic location of piRNA clusters among mammals,
although the precise sequence of each piRNA shows no apparent similarity [42,43,62]. This indicates that the relative
chromosomal position has some marked features with regard to production of piRNAs, and such special features aremaintained across mammals.
Caenorhabditis elegans has two PIWI proteins, PRG-1
and PRG-2. PRG-1 is required in germline maintenance,
and interacts with a class of small RNAs, called 21URNAs [63,64]. By definition, 21U-RNAs are the piRNAsof C. elegans. As their name implies, they are characterized by a first U bias, and their length is exclusively
21 nt, which is shorter than that of piRNA species inother organisms [65]. The vast majority of the 21URNAs are derived from thousands of individual locibroadly scattered in two large clusters on chromosome
IV [65]. These regions are gene-poor compared withother regions of the genome. A marked feature of 21URNAs is the existence of a clear cis motif located around40 bp upstream of the 21U-RNA encoding site [65]. The
consensus motif is CTGTTTCA and is flanked by an
AT-rich sequence, which is specifically recognized by
Forkhead family transcription factors [65,66]. In addition,
ChIP-on-chip experiments have shown a low level ofhistone H3 across the two piRNA clusters, which correlates well with DNase-sensitive sites [66,67]. Moreover, it
was also revealed that each upstream consensus motif
corresponds with the nucleosome-depleted region (NDR)[66]. These findings strongly suggest that each piRNA in
C. elegans is produced from an independent transcription unit.
Tetrahymena thermophila has a unique genome processing mechanism, called ‘programmed DNA elimination’. Most ciliated protozoans, including T. thermophila,
exhibit nuclear dimorphism, with a germline micronucleus (Mic) and somatic macronucleus (Mac) [68]. The
genomic sequence of this organism is processed during
the course of meiosis. Mic has an unprocessed genome,
and Mac has a processed one, but has a much larger genome size due to polyploidy. In contrast to the role of Mic
as a reservoir of genetic information, gene expression forPage 6 of 12

maintaining the organism takes place in Mac. The smaller
genome size of Mac compared with Mic is attributable to
DNA elimination induced by scan RNA (scnRNA). Internal eliminated sequences (IESs) are specific regions thatare selectively eliminated from the developing Mac genome, and there are over 6,000 IESs within the Mic genome.
scnRNA are loaded onto Twi, one of the Tetrahymena
PIWI proteins and are, therefore, T. thermophila piRNAs[69]. Twi1-scnRNA complexes are then transported to thedeveloping Mac, which has an unprocessed genome, andthey recognize and eliminate IESs through base-pairing between IESs and scnRNAs [70]. Strikingly, scnRNA production requires a Dicer-like protein, which is in clear contrast
to piRNA production in other animals [71]. scnRNAs map
predominantly to IESs, therefore, it can be said that IESs
are piRNA clusters in Tetrahymena [72]. Recent highthroughput analysis has uncovered biased transcription of
IESs in Mic; that is, IESs are destined to have high transcription activity [72]. Because of the lack of clear consensus sequence between different IESs, IESs are thought tobe epigenetically marked as piRNA clusters.
These findings in various animals suggest possible requirements to establish piRNA clusters, which are as
follows (in random order): 1) an ability to recruit
chromatin-modifying enzymes that contribute to the maintenance of open chromatin so as to attract and trap TEs, 2)an ability to recruit DNA specific factors (for example, specific transcriptional factors) to drive transcription of that
region, and 3) an ability to distinguish transcripts from that
region from other cellular transcripts and to specificallyprocess them into small RNAs (Figure 6B).
Transposition and chromatin boundaries

A prerequisite for genomic regions to act as TE traps isthat they must be frequent and non-deleterious sites for
TE insertion. TEs jump around the genome by transposition, but this appears to occur in a non-random manner[73]. The P-element is a DNA transposon that has beenused for insertional mutagenesis to isolate specific alleles
in Drosophila [74,75]. Because of this, a large body of data
has accumulated concerning the preferential P-element
insertion sites in the genome. Analysis of 100,000
transposition events identified that P-element insertion
preferentially occurs immediately 5′ to genes or within
5′ exons [76]. piggyBac, another TE that is also often
used for mutagenesis, also shows a high preference ofinsertion at or near promoter regions of genes [77].
These results indicate that these TEs tend to target
genomic regions that presumably contain open chromatin and/or are actively transcribed at the time oftransposition.
A fission yeast TE termed Tf1 is a retrotransposon
prevailing in the specific yeast genome. Tf1 insertion
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Figure 6 Model of Piwi-interacting RNA (piRNA) cluster formation. (A) Proto-piRNA cluster: transcripts are produced from a proto-piRNA-producing
locus. (B) Conversion to piRNA-producing locus: a specific transcription factor, histone marker, DNA methylation pattern, and/or RNA-binding protein (bluearrow, circle, and oval, respectively) convert the proto-piRNA-producing locus into a piRNA-producing site. (C) Sequential transposition event: the open natureof chromatin at the piRNA-producing locus attracts transposon integration (left panel). Certain types of transposons can accept the transposition
within themselves (right panel). (D) Maturation of piRNA cluster: a mature piRNA cluster is produced through sequential transposition eventsat piRNA-producing loci.

regions known to have relatively open chromatin [78,79].
These studies clearly argue for the relationships between
open chromatin and preferential transposition sites. However, it should be noted that these TE insertions at or nearpromoters alter the transcriptional activity of genes andare, therefore, often highly deleterious to the host. Thus,
individual genomes with these insertions tend to be eliminated from the population. So are there any genomic regions where TE insertions are tolerated?
In addition to gene promoters and their neighboring regions, chromatin boundaries are also known to have relatively open chromatin structures. A chromatin boundary
can act as a buffer between two functional chromatin domains by resisting the proliferation of epigenetic changes
that are characteristic of each, thus genes present in one
domain are not affected by regulatory sequences present
in a different domain [80-84] (Figure 7). Cis-regulatory elements are located at chromatin boundaries, and have different compositions of trans-acting proteins. They limit
the spreading of heterochromatin domains into regions ofactively transcribed genes (and vice versa) and prevent adventitious interactions between enhancers and promoterswhen located between them (acting as “insulators”)[83,84] (Figure 7A). However, chromatin boundaries,
especially those in Drosophila, between constitutive
heterochromatin and euchromatin are not fixed but
stochastic, as evident in position effect variegation(PEV), in which the heritable inactivating influence ofthe heterochromatin on a neighboring gene can spreadin some, but not all, cells of the same cell type [85].

In fission yeast, tRNA gene clusters near to the site ofconstitutive heterochromatin, such as those aroundcentromere, serve as strong boundary elements that inhibit the encroachment of heterochromatin into the euchromatic region [86,87] (Figure 7B). One explanation ofthis phenomenon is that the high transcriptional activity
from tRNA genes creates a discontinuity in arrayed nucleosomes that serves as a barrier to the propagation ofheterochromatin [88,89]. This high transcriptional activity might also function by promoting the activity ofhistone-modifying enzymes that contribute to the maintenance of open chromatin conformation [90]. A number of chromatin boundaries are associated with active
promoters. In addition, the recruitment of histone acetyltransferase activity correlates well with barrier activity
in multiple organisms [82]. These results suggest thepossibility that some promoters or transcription units
with specific characteristics may determine their own
chromosomal environment to ensure their activity, thereby
allowing them to effectively resist and even counteract heterochromatin formation, probably by manipulating histone
modifications.
In addition to histone modifications, replacement ofcore histones with their variants appears to contributeto boundary formation. The ENCODE project revealed
that specific histone variants are highly abundant at
chromatin boundaries. For example, H2A.Z is an evolutionarily conserved H2A variant present in all eukaryotes[91], which exhibits a characteristic localization in genomes, with high concentrations at gene promoters,

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Figure 7 Three types of boundary elements. (A) Boundary element intercepts the effect of an enhancer to the nearby promoter. (B) Boundary
element between heterochromatin and euchromatin serves as a barrier against the spreading of heterochromatin. (C) Boundary elements
residing in the BX-C region regulate the three homeotic genes to ensure the correct level and pattern of expression, thereby making possible
proper segmentation in the Drosophila embryo.

enhancers, and chromatin boundaries [17,92-95]. These
H2A.Z–rich regions are common NDRs, and are therefore DNase-hypersensitive. H2A.Z, together with H3.3, ahistone H3 variant, forms histone octamers, which constitute the most labile nucleosome state in human cells.
This leads to the dissociation of nucleosomes from chromatin, thereby forming NDRs [93,96]. Mapping the preferential H3.3 deposition sites in Drosophila S2 cells
revealed that there are specific sites at which H3.3 is heavily deposited [97,98]. The bithorax complex (BX-C) regulates the identity of each of the segments that contributes
to the posterior two-thirds of the fly [99]. The region encodes three genes, Ultrabithorax (Ubx), abdominal A(abd-A), and Abdominal B (Abd-B). It has been shownthat nine body segments are defined by the combinationof expression level of the three genes. Boundary elements
demarcate the BX-C region into nine parts, making possible the differential expression pattern of the three genes.
Interestingly, the preferential deposition sites of H3.3
match well with the BX-C boundary elements, such as
Fab-7, Fab-8, and Mcp [98]. Moreover, those sites are independently identified as DNase-hypersensitive sites [100](Figure 7C). Therefore, both H2A.Z and H3.3 serve as

molecular indicators of open chromatin conformation.
Interestingly, both H2A.Z and H3.3 have been recoveredfrom genome-wide RNAi screening to identify factors required for transposon silencing in Drosophila [35]. Thus,
it is tempting to speculate that both histone variants are
involved in piRNA production, possibly through maintaining the boundary nature of piRNA clusters (see below).
Of note, certain types of TEs themselves also showhigh rates of H3.3 deposition [97], implying that a TE itself can be a good recipient of a transposon. In addition,
it is known that transposition of retrotransposons tends
to occur within even older retrotransposons. For example,
nearly all retrotransposon insertions in the Arabidopsis
genome are into older retrotransposons [101,102]. The
recent ENCODE project has also revealed that DNase I
hypersensitive sites are strongly enriched at specific LTR
retrotransposons in the human genome in some cultured
cells, suggesting the possibility that TEs can transpose into
certain types of TE [95]. This would explain the reason
why TEs in piRNA clusters, such as flamenco, tend to bearranged in a nested fashion.
Together, these findings suggest that the relatively
open nature of chromatin at the chromatin boundaryYamanaka et al. Mobile DNA 2014, 5:22
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makes this region a susceptible site for TE transposition.
We propose a model in which the insertion of a single
TE in the chromatin boundary may trigger a runaway
process [103]; once the first TE inserts into the region,
this site becomes a stretch of landing pads for new TEs,
without deleterious consequences. Thus, in effect, anyslight concentration of TEs in a chromatin boundary
seeds a local TE expansion to produce an even more
preferential site or trap for transposition, creating an island or cluster of TEs (Figure 6C, D). It is well known
that the gypsy retrotransposon serves as an enhancerblocking insulator, a type of boundary element, wheninserted between promoter and enhancer [104]. Therefore, this gypsy insulator locus could be a prototype for
TE transposition landing pads. The aforementioned findings in Drosophila, mouse and other animals also imply
that special chromatin status with accompanying transcriptional factors and/or epigenetic factors at the chromatin boundary can give transcriptional license to that
region [22,61,66,72]. There is increasing evidence that
TEs often carry with them an array of transcription factor binding sites that, when integrated into the genome,
can become either alternative promoters or new enhancers [105]. Thus, transposition to a chromatin boundary of a TE that has a specific transcription factor binding
site, the transcription factor for which is already expressedin gonads, may make that region transcriptionally active
and put it under the control of the transcription factor. In
this way, boundary-specific elements may drive transcription of that boundary region to produce transcripts in gonads. A study describing the relationships between TE
insertion and de novo piRNA production shows that not
all TE insertions drive de novo piRNA production [106].
The transcriptional status at the insertion site might affectwhether the TE transcript is processed into piRNA [106].
This is consistent with the view we have discussed. The
chromatin boundaries are gene-poor regions, and therefore TE transposition at those regions is likely to be
neutral to the host, thereby allowing not only TE accumulation at those regions, but accumulation of nucleotidechanges in those accumulated TEs. Repeated transposition
events at the same boundary region would expand the sizeof clusters. Thus, it is possible that special transcriptional
units in the boundary regions are primitive piRNA production sites.

What makes the piRNA cluster so special?

When thinking about the process by which piRNA clusters are formed, the biggest outstanding question is how
does a specific locus turn into a piRNA-producing site?
In other words, what is the prerequisite for certain locito produce piRNAs? We propose two scenarios basedon the data described so far.

Page 9 of 12

One model is that piRNA production loci are marked
by specific factors. The very recent study from the Theurkauf laboratory revealed that dual-strand transcription
and recruitment of Rhino to the corresponding loci trigger
piRNA production [107]. Moreover, a study from theBrennecke laboratory showed that Rhino recruits Cutoff,which possibly acts to suppress transcription termination[55]. This implies that Rhino helps Cutoff and other additional factors to recognize nascent transcripts frompiRNA clusters, and to distinguish them from othertranscripts.
Another model is that transcripts from piRNA clusters
have some special property allowing them to be processed into piRNA, and this property is used by thepiRNA-producing machinery to distinguish piRNA transcripts from the vast majority of other transcripts. This
special property can be either altered splicing, characteristic 3′-end processing, or specific cis elements that direct recognition by special trans factors. Recently, Madhani
and colleagues showed that stalled spliceosomes are a signal for an RNAi response in a human pathogenic yeast,
Cryptococcus neoformans [108]. These authors proposedthat splicing intermediates are a preferred substrate forsmall interfering RNA biogenesis. This work explains how
specific transcripts are differentially recognized by thesmall RNA biogenesis machinery. It was recently reported
that Rhino can suppress normal splicing in the Drosophila
germ line with the aid of Uap56, making the piRNA precursor transcript different from other pol II transcripts[55,107,109]. However, in Drosophila follicle cells, splicing
of a long single-stranded transcript (more than 150 kb)
produced from the flam locus was reported [54]. Furthermore, the first intron of flam was found to be constitutively spliced [54]. In addition, there are numerous 3′-end
processing signals of TEs located in the flam locus. Therefore, there could be a certain mechanism that suppresses
transcription termination and poly(A) addition for theflam transcripts. Therefore, the transcript itself is sending
some message that it is different from other transcript.

Conclusions
Recent genome-wide ChIP analyses have revealed the locations on the genome where specific transcription andepigenetic factors sit. Cross-linking immunoprecipitation(CLIP) methods have also revealed specific binding sites
on transcripts for RNA-binding proteins. There is no
doubt that these types of analysis will propel this fieldforward and expand our knowledge of how piRNA clusters are formed and how transcripts from the clusters
are specifically processed into piRNAs. In addition, othermethods that are complementary to ChIP and CLIP
should also be applied to piRNA research. For example,
we do not have a comprehensive understanding of therepertoire of proteins that bind to piRNA clusters or toYamanaka et al. Mobile DNA 2014, 5:22
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the transcript from piRNA cluster. Taking advantage of
specific DNA-protein interactions, such as LexA with
LexA-binding sites, LacI with LacO repeats and modified
transcription activator-like effector (TALE), recent studies
have successfully immunopurified a chromatin locus ofinterest and identified associated proteins [110-113]. A
combination of RNA-binding proteins and their specificbinding sites, such as MS2 and BoxB sites, can be applied
to identify the proteins that bind to piRNA transcripts.
These types of strategy will allow us to identify the hidden
triggers for piRNA production.
Abbreviations
ChIP: Chromatin immunoprecipitation; CLIP: Cross-linking
immunoprecipitation; IES: Internal eliminated sequence; LINE: Long
interspersed repetitive element; miRNA: microRNA; NDR: Nucleosome-depleted
region; Nt: Nucleotide; PEV: Position effect variegation; piRNA: Piwi-interacting
RNA; RISC: RNA-induced silencing complex; scnRNA: Scan RNA; SINE: Shortinterspersed repetitive element; siRNA: Small interfering RNA;
TALE: Transcription activator-like effector; TE: Transposable element;tRNA: Transfer RNA.
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
SY and HS designed the structure of the review, and wrote the paper alongwith MCS. All authors commented on the manuscript, and read andapproved the final manuscript.
Acknowledgements
We thank Yasunori Aizawa, Kojiro Ishii, and Yota Murakami for critical readingof the manuscript. We are grateful to the members of the Siomi Laboratoryfor discussions. This work was supported by MEXT grants to HS, MCS, and SY,
and a Keio University Grant-in-Aid for Encouragement of Young Medical
Scientists to SY.
Author details
1
Department of Molecular Biology, Keio University School of Medicine,
35 Shinanomachi Shinjuku-ku, Tokyo 160-8582, Japan. 2 Department of
Biological Sciences, Graduate School of Science, The University of Tokyo,
Tokyo 113-0032, Japan.
Received: 18 June 2014 Accepted: 9 July 2014
Published: 1 August 2014
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doi:10.1186/1759-8753-5-22
Cite this article as: Yamanaka et al.: piRNA clusters and open chromatin
structure. Mobile DNA 2014 5:22.

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