Small non-coding RNAs in animal development

Abstract

The modulation of gene expression by small non-coding RNAs is a recently discovered level of gene regulation in animals and plants. In particular, microRNAs (miRNAs) and Piwi-interacting RNAs (piRNAs) have been implicated in various aspects of animal development, such as neuronal, muscle and germline development. During the past year, an improved understanding of the biological functions of small non-coding RNAs has been fostered by the analysis of genetic deletions of individual miRNAs in mammals. These studies show that miRNAs are key regulators of animal development and are potential human disease loci.

Full text

The traditional view of gene expression has relegated RNA
to a somehow subsidiary role, reserving the main regu-
latory functions for proteins. Nonetheless, as early as 1961,
Jacob and Monod proposed that RNAs could inhibit the
expression of operons by base-pairing with the operator
sequence
1
. The range of functions attributed to RNA has
substantially expanded with the realization that RNA func-
tions in the catalysis of crucial cellular processes, includ-
ing pre-mRNA splicing and protein synthesis. In the past
15 years, the discovery of RNA-silencing phenomena that
are mediated by an expanding assortment of small, non-
coding RNAs has unveiled the ability of RNA to impact on
an unanticipated variety of biological processes through
the post-transcriptional modulation of gene expression.
In particular, microRNAs (miRNAs) and
Piwi-interacting
RNAs (piRNAs) are implicated in various aspects of animal
development (for example, neuronal, muscle and germline
development). These recent advances place miRNAs and
piRNAs firmly on the map of key developmental genes
and point to their involvement in human diseases such as
birth defects and cancer (reviewed in REF. 2).
From their humble discovery as regulators of develop-
mental timing in Caenorhabditis elegans
3,4
, miRNAs have
emerged as important regulators of development in multi-
ple plant and animal species. The first miRNAs described
in animals, lin-4 and let-7, were identified by forward
genetics as controllers of the timing of larval develop-
ment in C. elegans: mutations of these genes resulted in
the reiteration of larval cell fates and retarded the final dif-
ferentiation of subsets of specialized cells
3–5
. Subsequent
studies in invertebrates have shown the widespread
involvement of miRNAs in various developmental pro-
cesses. In C. elegans, miR-61 and miR-84 have been shown
to modulate the expression of two orthologues of human
oncogenes, vav and ras, in the context of development of
the vulva
6,7
. Furthermore, lsy-6 and miR-273 are involved
in a complex gene regulatory network that establishes the
left–right asymmetry in the ASE chemosensory neurons
(reviewed in REF. 8). In Drosophila melanogaster, the range
of known functions of miRNAs is also wide
9,10
. The two
miRNAs that are encoded by the iab‑4 locus (which
are homologues of vertebrate miR-196) control expres
-
sion of the Ultrabithorax gene and induce the homeotic
transformation of halteres to wings when these miRNAs
are ectopically expressed
11
. Bantam, miR-2, miR-6 and
miR-14
regulate tissue growth through modulation of
both apoptosis and cell proliferation
9,10,12–14
. Important sig-
nalling pathways, such as the Notch and epidermal growth
factor pathways, are under the control of miRNAs
9,15,16
,
while the response to the steroid hormone ecdysone is
modulated by miR-14 through regulated expression of the
ecdysone receptor
17
.
The genomic distribution of miRNAs in invertebrates,
and to an even greater extent in vertebrates, is characterized
by the presence of families of several identical or closely
related mature miRNAs, which are sometimes encoded
within the same genomic cluster (reviewed in REF. 18).
A degree of functional redundancy among miRNAs is
therefore to be expected, and studies of C. elegans strains
that carry mutations of multiple miRNA-encoding loci
suggest that this is indeed the case in certain instances
19,152
.
However, several studies in the past year have shown that
the deletion of single miRNAs can result in discernable
phenotypes in mammals. Here, we review recent exciting
discoveries that show that miRNAs regulate important
developmental processes in vertebrates. We also include
Department of Molecular,
Cellular and Developmental
Biology, Yale University,
266 Whitney Avenue,
New Haven, Connecticut
06520, USA.
e-mails:
giovanni.stefani@yale.edu;
frank.slack@yale.edu
doi:10.1038/nrm2347
Published online
13 February 2008
Piwi-interacting RNAs
(piRNAs). Short RNA molecules
(24–30 nt long) that are
processed in a Dicer- and
Drosha-independent manner.
They associate with Piwi
proteins and have a role in
transposon silencing in flies.
In mammals, they are restricted
mostly to male germ cells.
Small non-coding RNAs in animal
development
Giovanni Stefani and Frank J. Slack
Abstract | The modulation of gene expression by small non-coding RNAs is a recently
discovered level of gene regulation in animals and plants. In particular, microRNAs (miRNAs)
and Piwi-interacting RNAs (piRNAs) have been implicated in various aspects of animal
development, such as neuronal, muscle and germline development. During the past year,
an improved understanding of the biological functions of small non-coding RNAs has
been fostered by the analysis of genetic deletions of individual miRNAs in mammals.
These studies show that miRNAs are key regulators of animal development and are
potential human disease loci.
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Nature Reviews | Molecular Cell Biology
m
7
G
m
7
G
AAAAAAAAAAAAAA
eIF4E
eIF4E
eIF4G
PABP PABP PABP PABP
AAAAAAAAAAAAAA
m
7
G
AAAAAAAAAAAAAA
eIF4E
eIF4G
PABP PABP
AGO
AGO
Ribosome
drop-off
Nascent
protein degradation
40S
60S
3′
5′
m
7
G
AAA
AGO
3′
5′
3′
5′
A
A
A
DCP1
DCP2
GW182
a Active translation
b
Inhibition of initiation
c Post-initiation
inhibition
d mRNA degradation
RNase III
One of a highly conserved
family of endoribonucleases
that cleave double-stranded
RNA and have an important
role in the maturation of
ribosomal RNA, among other
processes.
an overview of the roles of piRNAs, a recently discovered
class of small non-coding RNA, in germline development.
Given the breadth of the field, we focus our attention on
the most recent literature (for excellent reviews based
on the small RNA literature up to 2007, see REFS 20,21).
miRNA-mediated gene regulation
miRNAs are a class of short (19–25 nucleotide (nt)), single-
stranded RNAs that are present in plants and animals
18
.
Although they were previously thought to be exclusively
present in multicellular organisms, miRNAs have recently
been described in Chlamydomonas reinhardtii, a unicellular
alga
22,23
. miRNAs can be encoded in independent transcrip-
tion units, in polycistronic clusters or within the introns
of protein-coding genes
18
. They are transcribed, mostly
by RNA polymerase II, as capped and polyadenylated
primary miRNAs (pri-miRNAs) that contain extended
hairpin structures. Pri-miRNAs are cleaved in the nucleus
by the RNase III enzyme Drosha, releasing the shorter
(~65 nt long) precursor miRNA (pre-miRNA) hairpin
structure (reviewed in REF. 24). Independently of Drosha,
a subset of pre-miRNA hairpins can also be generated
Box 1 | Mechanisms of miRNA-mediated gene regulation
The mechanism of microRNA (miRNA)-mediated gene regulation, and how it affects active translation (see figure, part a),
is a matter of controversy. In essence, the available data support two possible views: first, the translation of mRNAs is
inhibited at the level of initiation, and the silenced mRNAs are occupied by few or no ribosomes
131
; second, the inhibition
takes place at a step that is subsequent to initiation, and the silenced mRNAs sediment in the polyribosome fractions in a
sucrose gradient. The former view has recently received support from several studies: Argonaute protein AGO2 has been
shown to bind the m
7
G cap of mRNAs through a domain that shares structural features with the translation initiation factor
eIF4E, suggesting that when AGO2 is recruited to the 3′ UTR of a target mRNA by miRNAs, it hinders the m
7
G cap
recognition by the translation apparatus
132
(see figure, part b). Consistently, in an in vitro system, increased levels of the
eIF4F complex (which includes the m
7
G cap-binding eIF4E translation factor) reversed miRNA translational inhibition
133
.
In further support of an effect on the translation initiation step, the assembly of the 48S complex (the translational complex
that precedes the addition of the large ribosomal subunit to form the competent ribosome) was found to be inhibited by
miR-2
in vitro, and eIF6, which inhibits joining of the 60S and 40S ribosomal subunits, was co-purified with the RNA-induced
silencing complex (RISC)
134,135
.
Nonetheless, mRNA that is inhibited by miRNAs has also been found to be associated with actively translating polysomes,
suggesting that, at least in a subset of cases, miRNA does not inhibit the initiation of translation
136–139
. The post-initiation
inhibition by miRNAs could result from rapid degradation of the protein product encoded by the targeted mRNA, or from a
high rate of ribosome drop-off during elongation, resulting in incomplete protein products that would be rapidly
degraded
138,139
(see figure, part c). Furthermore, the observation of translationally repressed mRNAs that co-sediment with
polysomes can be explained by the formation of dense, translationally silent mRNPs (‘pseudo-polysomes’)
134
.
A further element of uncertainty about the mechanism of action of miRNAs derives from observed variable levels of target
mRNA degradation, and colocalization of the RISC component with mRNA degradation factors in the P bodies (reviewed in
REF. 140; see figure, part d). Sequestration of mRNAs in the P bodies and degradation could be a step that follows blocking
of translation, or a causative event in miRNA repression. GW182, a P-body component, has recently been shown to interact
directly with AGO protein and to be recruited to the target mRNA in a let-7-dependent manner
141,142
. DCP1/2, decapping
protein-1/2; eIF4G, eukaryotic initiation factor-4G; GW182, a conserved member of the GW182 protein family that is
crucial for miRNA-mediated gene silencing; PABP, poly(A)-binding protein.
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miRNA-induced silencing
complex
(miRISC). A multicomponent
gene regulatory complex that
is activated by a microRNA
(miRNA) associated with an
Argonaute protein and that
regulates gene expression,
mediated by the sequence
complementarity between the
miRNA and the target mRNA.
Argonaute protein
One of a family of
evolutionarily conserved
proteins that are characterized
by the presence of two
homology domains (PAZ and
PIWI). Argonaute proteins are
essential for diverse RNA-
silencing pathways.
P bodies
Cytoplasmic foci that are
thought to store and degrade
translationally repressed RNA.
from introns by the combined actions of the spliceosome
and the lariat-debranching enzyme (LDBR)
25–27
. On export
into the cytoplasm by exportin-5, the pre-miRNA is fur
-
ther processed by a second RNase III, Dicer, which excises
a 19–25-nt double-stranded duplex. This short duplex is
incorporated into the functional miRNA-induced silencing
complex (miRISC), where the mature miRNA strand is
preferentially retained. The miRISC contains miRNA, an
Argonaute protein and other protein factors and is the effector
complex of the miRNA pathway.
The miRISC is directed to mRNAs that are comple
-
mentary to its miRNA component. miRISC inhibits the
expression of mRNAs in one of two ways, depending on
the degree of complementarity between miRNA and the
target. If the complementarity is perfect, as is mostly the
case in plants, the target mRNA is cleaved and degraded.
By contrast, the complementarity between miRNAs and
their targets in animals is frequently imperfect, and the
mechanism leading to inhibition of mRNA expression
is not well understood. Various mechanisms have been
documented, including translational inhibition at the
level of initiation and elongation, rapid degradation of
the nascent peptide, mRNA sequestration into P bodies
and mRNA degradation (reviewed in REF. 28). It is likely
that features of the mRNA or of the proteins bound to it
determine the method of suppression; however, the key
features and methods of repression remain an intense
focus of current research (BOX 1).
As noted above, animal miRNAs bind with imperfect
complementarity to their targets, resulting in a variable
degree of miRNA–target mismatches. As such, the search
for targets of miRNAs is not straightforward in animals.
Many studies have underscored the importance of high
complementarity between residues 2–8 at the 5′ end of
the miRNA with its target site, referred to as the seed
region
29,30
. This model has recently been refined to account
for the presence of secondary structures and other features
of the 3′ untranslated region (UTR) sequence surrounding
the target site, and for the ability of complementarity at the
3′ end of the cognate miRNA to compensate for imperfect
seed matching
31–34
.
The impact of miRNA-mediated biological regulation
is estimated to be vast: hundreds of miRNAs have been
cloned and thousands more have been predicted bio-
informatically
35–38
. Furthermore, experiments in vitro
have shown that overexpression of single miRNAs can
result in decreased levels of >100 mRNAs, leading to the
hypothesis that a large fraction of protein-coding genes are
regulated by miRNAs
39
. Global analysis of mRNA levels
relative to miRNAs shows low or undetectable levels of
expression of predicted target mRNAs in tissues that
express the targeting miRNA
40,41
. These observations have
been interpreted as indicating that one role of miRNAs
may be to function as developmental switches or, more
subtly, to sharpen the borders of spatial or temporal gene-
expression domains. Here they would ensure the silencing
of unwanted messages resulting from leaky transcription
or previous synthesis (reviewed in REFS 20,42), or allow
the maintenance of target mRNA expression within an
optimal range
43
. Therefore, it has been proposed that
one function of miRNA-mediated control of gene expres
-
sion in vertebrates may lie in conferring robustness to
developmental programmes
42,20
.
The global functional role of miRNAs in development
can be inferred from animals that lack Dicer1 or DGCR8,
a cofactor that is required for Drosha function (TABLE 1).
Deletion of Dicer1 and Dgcr8 results in early developmen-
tal arrest in mice, accompanied by defects in the prolifera-
tion of pluripotent stem cells
44–46
. Tissue-specific deletion
of Dicer1 in mice and ablation of dicer in zebrafish results
in a seemingly unaffected overall patterning, while the
establishment, maintenance and function of subsets of
cells are impaired to variable degrees
47–53
.
The role of individual miRNAs in the development
of mammals has only recently begun to be assessed by
genetic ablation. Next, we summarize recent studies that
relate the functions of individual miRNAs to the regula-
tion of early embryonic development or to the subsequent
development and function of various tissues.
miRNAs in early embryonic development
Several recent studies have revealed a substantially con-
served network of intercellular signalling mechanisms that
specify the site of initiation of gastrulation in vertebrates
and, therefore, the embryonic axis (reviewed in REF. 54).
Table 1 | Mouse deletions of RNA-encoding and RNA-silencing genes
Gene deleted Phenotype Refs
Mir-1-2 Cardiac morphogenetic defects, cardiac electrophysio-
logical defects, fatal with variable penetrance.
52
Mir-208 Absence of cardiac hypertrophy in stress conditions,
failure to upregulate βMHC in stress conditions.
82
Mir-155 Defective adaptive immunity, fibrosis and infiltration of
the lung, defects in germinal centre reaction (decreased
interleukin-2, interferon-γ), decreased production of
immunoglobulin.
101, 102
Mir-150 Expanded lymphocyte B1 population, decreased
lymphocyte B2, increased immunoglobulin production,
increased c-Myb.
144
Miwi2 Male sterility, spermatogenesis arrest at early prophase
meiosis I, complete loss of spermatogonia in adults.
122
Miwi Male sterility: spermatogenesis arrest at early round
spermatids stage.
120
Mili Male sterility: spermatogenesis arrest at early prophase
meiosis I.
121
Dicer-1 Lethality in early embryonic stages, depletion of
multipotent stem cells, embryonic stem cells unable to
differentiate, loss of epigenetic silencing of centromeric
sequences.
44,45
Dicer-1
(tissue-
restricted
deletion)
Limb morphogenesis defects, lung development
defects, incomplete embryonic myogenesis, loss of
Purkinje cells in adult cerebellum, evagination of
hair germs, epidermis hyperproliferation, impaired
development of αβ-expressing thymocytes,
reduced development of ventricular myocardium.
51,
145–149
Ago2 Lethality at mid-gestation, lethality in early embryonic
stages.
150,151
Dgcr8 Lethality in early embryonic stages, embryonic stem
cells unable to differentiate.
46
Ago2, Argonaute-2; βMHC, β-myosin heavy chain; c-Myb, transcription factor and proto-
oncogene; Dgcr8, Drosha cofactor; Mili, Piwi-like homologue-2; Miwi, Piwi-like homologue-1;
Miwi2, Piwi-like homologue-4.
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In the Xenopus laevis embryo, β-catenin accumulates in
the future dorsal side in response to fertilization and syner-
gistically interacts with Vg-1 to induce the Nieuwkoop
centre, which induces the Spemann’s organizer, which, in
turn, initiates gastrulation and induces the ectoderm to
become the neural plate and neural tube. The transform-
ing growth factor-
β (TGFβ) protein Nodal has a pivotal
role in the induction and patterning of mesoderm: a
dorsal-to-ventral gradient of Nodal activity is required
for the formation of dorsal mesoderm and Spemann’s
organizer
55,56
.
The mechanism that translates the early β-catenin
dorsal–ventral gradient into the late blastula gradient of
Nodal activity is not known. A recent study shows that,
at least in X. laevis, miRNAs have a relevant role in this
process
57
. Two miRNAs, miR-15 and miR-16, inhibited
Nodal signalling by reducing the expression of one of
its receptors, Acvr2a. Consistently, Spemann’s organizer
and head structures were reduced by overexpression
of miR-15 and miR-16, but were increased by blockage of
these miRNAs. Furthermore, miR-15 and miR-16 are
epistatic to β-catenin, as blockage of miR-15 and miR-16
restored dorsal mesoderm induction in embryos in
which Wnt/β-catenin signalling was suppressed. These
data, and the observation of a ventral-to-dorsal gradient
of miR-15 and miR-16 that is reciprocal to the
β-catenin
gradient, suggest that inhibition of miR-15 and miR-16
expression is a major mechanism through which the Wnt
signalling pathway promotes Nodal signalling and dorsal
mesoderm patterning
57
.
Evidence of a role for miRNAs in the modulation
of Nodal in the early embryo is seemingly at odds with
the observed absence of gross defects of embryonic axis
specification in zebrafish that lack Dicer
47
. A possible
explanation for this discrepancy is provided by the
absence of potential complementary sites for miR-15
and miR-16 in the 3
′ UTR of zebrafish Acvr2a, whereas
such sites are present in mammals, which require Dicer
for early embryonic development
57,44
. However, other
components of the Nodal signalling pathway in zebrafish
are under the control of miRNA-mediated modulation.
miR-430, a highly abundant miRNA that is required for
the clearance of maternal mRNAs, has recently been
shown to directly decrease the expression of squint
(sqt), a member of the Nodal family
58
. Interestingly, lefty,
an antagonist of Nodal, is also regulated by miR-430.
The simultaneous relief from miR-430-mediated regula
-
tion of both squint and lefty resulted in either a modest
effect or no effect on mesoderm induction, whereas
other outputs of Nodal activity (such as the number of
endoderm progenitors and specialized dorsal forerunner
cells) were decreased.
Therefore, it was suggested that miR-430 fine-tunes
the overall activity of the Nodal signalling pathway by
balancing the relative levels of agonist and antagonist.
Furthermore, miR-430 was shown to confer robustness
by dampening the levels of signalling molecules because
overexpression of squint or lefty did not produce an
appreciable phenotype in the presence of miR-430, but
resulted in disruption of development when miR-430
complementary sites were mutated
58
.
miRNAs in neuronal development
It has long been suggested that the nervous system, with
its astonishing variety of functionally specialized cellular
subtypes and vast number of synaptic contacts, requires
ways to expand the information content of a limited
number of protein-coding genes more than any other
tissue. A well-documented means of achieving this goal
is alternative splicing (reviewed in REFS 59,60). Small
non-coding RNAs offer another source of complexity.
The ability of miRNAs to specify and maintain neuro-
nal cell-type identity is strikingly demonstrated by the
requirement for lsy-6 and miR-273 in the establishment
of left–right asymmetry in the ASE neurons in C. elegans
(reviewed in REF. 8). The role of miRNAs in late neuronal
development, neuronal functions and synaptic plasticity
have been exhaustively reviewed elsewhere
61
. In addition,
recent evidence points to a role for miRNAs in neuronal
cell differentiation. For instance, the neuronal-tissue-
specific miR-124 helps to acquire and maintain the neuro-
nal cellular identity by directly silencing a large number
of target mRNAs, and through the repression of master
regulators of gene expression.
miR-124 is expressed specifically and abundantly in
the mouse brain and in P19 pluripotent cells on their
differentiation to neuron-like cells
62,63
. Mis-expression of
miR-124 in HeLa cells inhibited the expression of >100
genes that are normally expressed at low levels in neuronal
tissue, suggesting that it may contribute to neuronal differ-
entiation
39
. One target of miR-124 is polypyrimidine tract-
binding protein (PTB, also called PTBP1 or hnRNP-I), a
regulator of alternative splicing that inhibits the inclusion
of alternative cassette exons
53,64
. During neuronal differ-
entiation, the switch between the expression of PTB and
nPTB (neuronal PTB, also called PTBP2 or brPTB), a
highly homologous neuron-specific protein encoded by
a separate gene, results in widespread changes in the splic-
ing pattern of genes that are involved in crucial neuronal
functions
65
. The mutually exclusive expression of the two
PTB forms is directly enforced by PTB, which alters the
splicing of nPTB by repressing the inclusion of an alter-
native exon, resulting in a message that carries a premature
stop codon
65,53
. Therefore, miR-124 indirectly activates the
expression of nPTB by inhibiting PTB
53
(FIG. 1a).
Consistent with the inhibition of PTB by miR-124, a
strikingly complementary pattern of expression of PTB,
nPTB and miR-124 was observed in mouse embryos. PTB
was expressed in areas of the developing neuronal system
where non-differentiated progenitor cells are present,
whereas nPTB and miR-124 were expressed in differenti
-
ated neurons. Furthermore, the distribution of the exon-
including isoforms of various genes regulated by PTB
and/or nPTB precisely overlapped with miR-124 expres
-
sion, which is consistent with the ability of miR-124 to
antagonize PTB. Finally, the pattern of expression of
splicing isoforms of PTB target genes was perturbed
in mice carrying a telencephalon-restricted
Dicer-null
mutation, confirming that miRNAs are involved in the
regulation of splicing
53
. The opposite switch between PTB
isoforms, from nPTB to PTB, is also regulated by miRNAs;
during muscle development, the muscle-specific miR-133
directly inhibits the expression of nPTB in myoblasts,
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Myoblasts Neuroblasts
Myotube
Muscle fibre Neurons
miR-133 nPTB nPTB
Progenitor
splicing
pattern
Neuronal-
specific
splicing
pattern
Mature
splicing
pattern
miR-124 PTB
a
b
resulting in changes in the splicing pattern of genes that are
regulated by nPTB
66
(FIG. 1b). Intriguingly, these are the first
described examples of an miRNA achieving a biological
effect by indirectly modulating alternative splicing.
Besides modulating the PTB–nPTB switch, miR-124
affects another crucial regulator of neuron-specific gene
expression, the RE1-silencing transcription factor (
REST,
also called NRSF). REST is a transcription factor that
represses the extra-neuronal transcription of several
genes, including miR-124 (
REF. 67, reviewed in REF. 68).
Reciprocally, miR-124 inhibits REST activity by targeting
small C-terminal domain phosphatase-1 (SCP1),
which
is required for REST-mediated repression of neuronal
genes
69
. These findings suggest the existence of a nega-
tive feedback loop: in non-neuronal cells and neuronal
progenitors, the expression of neuronal-specific genes
(including miR-124) is repressed by REST and SCP1;
as cells progress towards neuronal differentiation and
REST is transcriptionally inhibited, miR-124 ensures the
fast cessation of the biological effects of REST by post-
transcriptionally inhibiting the expression of its required
cofactor SCP1
(REF. 69).
miRNAs in muscle development
The formation of mature muscle proceeds with the
exit of myoblasts from the cell cycle, the expression of
muscle-specific genes and the suppression of genes that
are specific to other cell lineages and tissues. A role for
miRNAs in this process was originally suggested by an
enrichment of specific miRNAs in myocytes
63
. Blocking
miRNA maturation specifically in the heart by deletion
of Dicer led to heart failure at embryonic stages and poor
development of the ventricular muscle
52
. The overall
architecture of the heart chambers was grossly normal, as
were molecular markers of early heart differentiation and
patterning
52
. These broad observations led to the study of
a particular miRNA, miR-1, for its role in controlling the
development of skeletal and heart muscle.
miR‑1 and the development of heart and muscle. miR-1
is highly expressed in skeletal and heart muscle across
species from D. melanogaster to humans
63,70–72
. In mice
and humans, miR-1 and its variant miR-206 are encoded
by three separate loci (MIR‑1‑1, MIR‑1‑2 and MIR‑206);
each of these loci co-expresses a closely linked gene,
called MIR‑133 (REF. 73). Consistent with a crucial role
for miR-1 in the proper establishment and maintenance
of muscular and cardiac tissue, its expression is regu-
lated by transcriptional master regulators of myogenesis:
MEF (myocyte-specific enhancer-binding factor) and
MYOD (myoblast determination protein-1) are required
for somitic expression of MIR‑1‑1 and MIR‑1‑2, respec-
tively, whereas serum response factor (SRF) is required
for cardiac expression of both
33
.
In an in vitro model of myoblast differentiation, the
expression of miR-1 was induced on growth in a differentia
-
tion medium, and coincided with the appearance of muscle-
specific molecular markers
73
. Overexpression of miR-1
in myoblasts promoted differentiation while reducing
cell proliferation
73
. A similar activity was observed in the
developing heart, where miR-1 overexpression reduced
cell proliferation, resulting in a thinner ventricular wall
33
.
Figure 1 | MicroRNAs in neuronal development. Polypyrimidine tract-binding protein (PTB) and its neuron-specific
homologue nPTB are regulators of gene expression at the interface between RNA silencing and splicing. a | In
undifferentiated cells, the ubiquitous splicing regulator PTB represses the expression of nPTB by affecting the pattern of
pre-mRNA splicing. As differentiation proceeds, miR-124 activates the expression of nPTB by inhibiting PTB. nPTB, in turn,
shifts the alternative splicing of an array of genes to a neuron-specific pattern. Furthermore, miR
-124 silences REST,
a transcriptional inhibitor of neuron-specific genes that is expressed outside the neural system. b | As myogenesis
progresses from the myoblast stage to the myotube stage, the level of the muscle-specific miR-133 increases. miR-133
inhibits the expression of nPTB, indirectly affecting the pattern of alternative splicing of several target genes.
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miR-1
miR-208
HAND2 IRX5
KCND2
Heart
morphogenesis
Electric
conduction
Stress
Hypothyroidism
T3
TRE βMHC
TRAP
THRAP1 TR
TRE αMHC
a
b
Hyperplasia
Enlargement of an organ
resulting from an increased
number of its cells.
Recent elegant work in mutant mice has demonstrated
that miR-1 is not solely an early regulator of the prolif
-
eration-to-differentiation switch in muscle, but also has
important roles in later cardiac functions
33,52
. Selective
ablation of Mir-1‑2, one of the two genes encoding miR-1
in the mouse genome, resulted in animals with enlarged
hearts due to thickened walls. Cardiomyocytes in the
Mir-1‑2 mutant mice failed to exit the cell cycle properly,
resulting in hyperplasia. These defects, along with the fail-
ure of ventricular septation, led to the prenatal or early
postnatal death of approximately half of the mutants.
Surviving adult Mir-1‑2 mutant animals displayed a com-
plex array of electrophysiological defects that resulted in
sudden death, similar to clinical observations in humans
with abnormal electrocardiograph traces
52
.
miR‑1 targets. Similar to all miRNAs, miR-1 could poten-
tially regulate many genes, and putative targets have been
identified through a combination of computational and
experimental approaches
39
. During looping, a vital stage of
heart morphogenesis, miR-1 controls the balance between
proliferation and differentiation of myocardiocytes
through translational inhibition of HAND2, which
encodes a transcription factor
33
(FIG. 2a). Furthermore,
several other genes that are involved in cell-cycle regu
-
lation or cardiac growth and differentiation have been
suggested as possible targets of miR-1
(REF. 52).
The electrophysiological effects of miR-1 seem, at least
in part, to be mediated by its control of the transcrip-
tion factor IRX5, which in turn inhibits the expression
of a gene that encodes a potassium channel, KCND2
(REF. 52) (FIG. 2a). KCND2 has an important role in cardiac
repolarization, which suggests a potential mechanism
for the observed disturbances of cardiac conduction in
Mir‑1‑2 mutant mice. Interestingly, increased miR-1
levels are observed both in human patients with coronary
artery disease and in animal models of heart ischaemia.
Overexpression of miR-1 favoured the appearance
of potentially fatal arrhythmias, whereas its blockage
through chemically modified antisense oligonucleotides
reduced their occurrence
74
.
The importance of miR-1 function in skeletal muscle
development is highlighted in the Texel sheep, a breed that
has been selected over centuries for its increased muscu-
larity. A recent study has identified a G→A transition in
the 3′ UTR of a myostatin gene that negatively regulates
muscle mass in these sheep
75
. This single nucleotide
polymorphism optimizes a recognition site for miR-1,
resulting in decreased levels of myostatin in Texel sheep
and increased muscle growth
75
.
Although it is clear that miR-1 activity is a vital com
-
ponent of muscle development and function, several
aspects of miR-1-mediated regulation remain unclear. For
example, the two copies of miR-1 in the mouse genome
are expressed in a similar, although not identical, spatial
and temporal pattern. However, they do not appear to
act redundantly because ablation of Mir-1‑2 alone causes
a profound phenotype
52
. Furthermore, both copies of
Mir-1 are transcribed as a primary transcript, which also
contains Mir-133. miR-133 is detected at high levels spe-
cifically in the muscle and heart but, contrary to miR-1,
overexpression of miR-133 causes increased proliferation
and decreased myocyte differentiation
63,73
. The presum-
ably coincident and simultaneous transcription of these
two miRNAs with opposite effects on muscle maturation
prompts the question of whether a post-transcriptional
mechanism exists that balances their actions in differ-
ent phases of development. In addition, targets of miR-1
and miR-133 might be differentially expressed at differ
-
ent phases of development. The functional interaction
between miR-1 and miR-133 and the mechanism that
balance their actions awaits further elucidation.
miRNAs involved in cardiac hypertrophy. The postnatal
heart responds to various stress signals, such as hyper-
tension or endocrine dysfunctions, with a hypertrophic
(enlargement) response. This enlargement stems from
an increase in cardiomyocyte volume, not proliferation.
Although cardiomyocyte hypertrophy probably provides
a functional advantage in its early phase, it is soon accom-
panied by deposition of fibrotic tissue and decreased con-
tractility, and ultimately results in heart failure. The various
signalling cascades that are implicated in hypertrophy
Figure 2 | MicroRNAs in cardiac development. a | miR-1
regulates cardiac morphogenesis by optimizing the level
of the HAND2 transcription factor. Electric conduction is
abnormal in mice that lack miR-1 as a consequence of
de-inhibition of IRX5, a homeodomain-containing
transcription factor that represses the expression of the
KCND2 potassium channel. b | In normal conditions in
wild-type animals, miR-208 maintains an optimal level of
the thyroid hormone receptor (TR) cascade activity by
acting on THRAP1 (thyroid hormone receptor-associated
protein complex 240 kDa component) in a negative
feedback loop. In transgenic mice that overexpress
miR-208, inhibition of the TR pathways allows aberrant
expression of β-myosin heavy chain (βMHC) in the adult.
Similarly, in conditions of stress or hypothyroidism,
decreased activity of the TR cascade leads to expression
of βMHC and hypertrophy. In the absence of miR-208 in
null mice, THRAP1 is de-repressed and baseline levels of
TR activity are abnormally high and resistant to inhibition
by stress signals. Therefore, Mir-208-null mice do not
express elevated levels of βMHC or undergo cardiac
hypertrophy in conditions of stress and hypothyroidism.
T3, tri-iodothyronine; TRE, T3 response element.
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Deep sequencing
techniques
Sequencing to high coverage,
where coverage (or depth)
corresponds to the average
number of times that a
nucleotide is sequenced.
activate a set of transcription factors that have early roles
in cardiac development (reviewed in REF. 76). Several
groups of researchers have identified a set of miRNAs
that have abnormal levels of expression in mouse and rat
hypertrophy models
77–81
. A subset of these miRNAs, when
overexpressed, conferred the morphologic features that
are typical of hypertrophy in primary cardiomyocytes.
Furthermore, a single miRNA, miR-195, is sufficient
to provoke heart dilative hypertrophy when it is over-
expressed in vivo in transgenic mice. Interestingly, miR-195
expression is also elevated in failing human hearts
77
.
Several miRNAs were also reduced in murine and
human hypertrophic hearts, including miR-133 and miR-1
(REFS 79,80). These two miRNAs, which have a crucial
role in heart development and function (see above),
also affect cardiac hypertrophy
79,80
. Although miR-1
and miR-133 seem to act in opposition during skeletal
muscle differentiation, they function cooperatively in the
context of cardiac hypertrophy. Decreased levels of either
miR-133 or miR-1 are sufficient to initiate a hypertrophic
phenotype
73,79,80
.
A hallmark of cardiac hypertrophy is the aberrant
postnatal activation of fetal genes. For example, β-myosin
heavy chain (βMHC) is aberrantly expressed during
hypertrophy, at the expense of the adult form, αMHC.
βMHC has lower ATPase activity than the adult form,
and thus its expression results in contractile dysfunction
in the adult heart. Intriguingly, the heart-specific miRNA
miR-208 is encoded within an
αMHC intron, which
suggests the possibility of miRNA-mediated regulation.
To explore this possibility in vivo, Van Rooij and collab-
orators deleted miR-208 by homologous recombination,
without affecting the levels of expression of αMHC.
The phenotype of untreated mutant animals was subtle,
with decreased contractility and expression of fast skel-
etal muscle-specific genes in the heart. In experimental
cardiac hypertrophy models, however, Mir‑208-null
animals failed to show heart hypertrophy and induction
of βMHC, unlike wild-type animals. Reciprocally, trans-
genic overexpression of miR-208 was sufficient to induce
robust expression of βMHC. These data hint at a role for
miR-208 in setting the threshold of induction of
βMHC in
response to stress and hypothyroidism
82
(FIG. 2b).
Thyroid hormone receptor (TR), in combination
with THRAP1 (thyroid hormone receptor-associated
protein complex 240 kDa component), directly represses
the expression of βMHC and promotes αMHC at the
transcriptional level (reviewed in REFS 76,83). miR-208
maintains an optimal level of TR activity by negatively con-
trolling the expression of THRAP1 in a negative feedback
loop (FIG. 2b). In wild-type animals, this miR-208-mediated
inhibition is not sufficient to allow expression of βMHC,
but a threefold increase of miR-208 in transgenic mice
resulted in robust induction of βMHC. In the absence of
miR-208, the threshold of inhibition of the TR cascade is
elevated beyond the ability of hyperthyroidism and stress
stimuli to overcome it. Thus, a negative feedback loop
has been revealed whereby the same locus that encodes
αMHC also produces miR-208, which, by regulating the
TR pathways, also modulates the expression of the two
MHC genes and the contractility of the heart
82
(FIG. 2b).
miRNAs in lymphocyte development
The haematopoietic system offers an ideal model for stud-
ies that correlate gene regulation with cell lineage specific-
ation owing to the ease of isolation and expansion in vitro
of precursor and intermediate staged cells. In addition,
there is a wealth of molecular markers that are specific
for various phases of differentiation. Here, we focus our
attention on recent studies that demonstrate the role
of miRNAs in lymphocyte maturation. Studies that exam-
ine miRNA function in myeloid lineage development and
macrophage function have been reviewed elsewhere
84,85
.
miR‑181. The miRNA content of the haematopoietic sys-
tem has recently been surveyed using DNA microarray
and deep sequencing techniques
86–88
. Characterization
of the composition of the miRNA repertoire of cells
at various stages of T-lymphocyte maturation showed
that only a handful of miRNAs displayed significantly
altered expression levels across T-lymphocyte develop
-
ment
88
. Nonetheless, dynamic changes in the miRNA
abundance during T-lymphocyte maturation define
a miRNA ‘signature’ that is specific for each stage
88
. In
particular, miR-181 is elevated at the double positive
(DP) stage, when thymocytes expressing both CD4 and
CD8 undergo positive and negative selection, suggesting
a role for miR-181 in this process.
miR-181 appears to increase the sensitivity of DP cells
to stimulation of the T
-cell receptor (TCR). TCR signal
-
ling within thymocytes must be strictly regulated because
it is responsible for selecting cells that will only strongly
interact with non-self ligands. During development in the
thymus, the TCR on DP cells must bind to the major histo
-
compatibility complex (MHC)–peptide complex with low
affinity in order to be selected for further development in
lymphocytes. The negative selection of strong binders at
this stage is required to eliminate cells that could induce
autoimmunity. However, once the naïve lymphocytes exit
the thymus, the TCR must bind MHC–peptide complexes
with high affinity for activation into a mature lymphocyte.
The modulation of T-cell responsiveness is therefore
crucial for the cellular outcomes at different phases of
differentiation. T lymphocytes that overexpress miR-181
display a stronger activation of the TCR signalling cascade
in response to low-affinity MHC–peptide complexes com
-
pared with untreated cells
89
. Blocking miR-181 in DP cells
suppresses both positive and negative selection. Because
wild-type DP thymocytes have a tenfold higher miR-181
abundance than their mature counterparts, it appears
that miR-181 is responsible for the intrinsic modulation
of cellular sensitivity to TCR activation. miR-181 sets a
lower threshold of TCR cascade activation in DP cells
than in mature lymphocytes by repressing the expression
of several phosphatases, resulting in increased levels of
activation of two TCR signalling molecules, LCK (lym
-
phocyte cell-specific protein-tyrosine kinase) and ERK
(extracellular signal-regulated kinase)
89
(FIG. 3a).
miR‑155. The BIC gene (B-cell integration cluster) was
originally identified as a site of frequent integration
of avian leukosis virus, leading to B-cell lymphoma
induction
90
. BIC encodes a ~1,700-nt polyadenylated
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miR-155
c-MAF
IL-4
Th2
B cell
Macrophages
DP cells
CD4
Th1
Immature T helper
CD8
Lymph node
Mature
T lymphocytes
DP
Germinal centre
reaction
• Self antigens
(thymus)
• Negative
selection
• Positive
selection
• Foreign antigens
(periphery)
• Immune
response
ba
miR-181 ( )
T-receptor
activation
threshold ( )
Small interfering RNA
(siRNA). Short double-stranded
RNA molecules (~21–23 nt)
that guide the cleavage and
degradation of RNA that is
complementary to one of its
strands.
Germ-line stem cells
(GSCs). Cells that have the
ability to self-renew and to
generate differentiated cells
that are restricted to the germ
cell lineage.
Primordial germ cells
Embryonic cells that give rise
to the germ cell lineage.
and spliced transcript that lacks a recognizable protein-
coding sequence. The gene is poorly conserved except
for a ~100-bp region
91
. Interest in BIC has recently been
reignited by the finding that the conserved region encodes
miR-155
(REF. 92). BIC/miR-155 expression is increased
in activated B and T cells, macrophages and dendritic
cells
93,94
. Elevated levels of miR-155 have also been found
in Burkitt lymphoma, Hodgkin lymphoma, other B-cell
lymphomas, breast carcinoma and lung carcinoma
95–100
.
Furthermore, high levels of miR-155 correlated with a
poor prognosis in lung cancer
100
.
More recently, two groups have used genetic deletion
of the Mir-155 gene in mice to investigate its function
in vivo
101,102
. The absence of miR-155 results in a complex
alteration of the immune response, as determined by
tests for B-cell, T-cell and dendritic-cell function and by a
failure to achieve protective immunity against a bacterial
pathogen
102
. Furthermore, Mir-155-null mutants showed
a lung histopathology that was reminiscent of human
autoimmune diseases, with diffuse fibrosis, increased
collagen deposition and immune cells in the bronchioli
102
.
In the context of generalized altered homeostasis of the
immune system, specific abnormalities of the lymphocytes
were identified. First, Mir-155-null mutants displayed an
altered equilibrium between the two classes of helper
T lymphocytes, Th1 and Th2. This balance appears to be
shifted in favour of Th2 in Mir-155-knockout mice. This
result is, at least in part, explained by loss of miR-155-
mediated inhibition of c-MAF, a transcription factor that
promotes the expression of interleukin-4 (IL-4), one of the
major outputs of Th2 cells (FIG. 3b). Second, in Mir-155-
null mice, B lymphocytes were decreased in the germi
-
nal centres, which are areas within lymph nodes where
B lymphocytes divide, differentiate to plasma cells and
start immunoglobulin production
101
(FIG. 3b). The pheno-
type of Mir-155-null mice demonstrates a complex role
for miR-155 in various aspects of the adaptive immune
response. Further analysis of these mutants will probably
also reveal roles for miR-155 in innate immunity because
miR-155 expression in macrophages has recently been
described
94
.
piRNAs and germline development
A large part of eukaryotic genomes is occupied by trans-
posons and retrotransposons — repetitive sequences that
have duplicated themselves many times and can move
into new locations. Similar to retroviruses, retrotrans-
posons propagate themselves by transcription into RNA
from their location in the genome, followed by reverse
transcription back to DNA and integration into a new
genomic location. Activation of transposable elements
in the germline leads to the transmission of an increased
copy number to the next generation. As such, trans-
posons are mainly active in the germline. Because active
transposable elements lead to genomic instability, the
impairment of genes that are responsible for transposon
control often results in sterility and other abnormalities
of the germline. A role for small interfering RNA (siRNA)
in the control of retrotransposon mobility has previously
been demonstrated in C. elegans
103,104
. Recent studies are
beginning to elucidate how mechanisms that are based
on RNA silencing have specialized to curb the activity of
selfish genetic elements, specifically in the germline, by
deploying a novel set of short non-coding RNAs.
Repeat‑associated RNA and germline development in flies.
Forward-genetic studies provided early indications of an
involvement of proteins, which were later recognized as
members of the Argonaute family, in germline develop-
ment
105
. The piwi gene (P-element induced wimpy testis)
was first identified as a mutation that impairs asymmetric
division in germ-line stem cells (GSCs), resulting in severe
defects in spermatogenesis and female sterility
105,106
. Also,
Aubergine (encoded by aub) was originally identified as
a mutation that leads to sterility
107
. Aubergine is required
for the formation of pole cells, from which primordial
germ cells originate
108
. Piwi, Aubergine and Argonaute-3
(encoded by Ago3) are germline-specific Argonaute pro-
teins in D. melanogaster. The other two members of the
Argonaute family in flies, AGO1 and AGO2, which are
expressed more abundantly in the soma, are involved in
miRNA- and siRNA-mediated RNA-silencing pathways,
respectively, which strongly suggests an involvement of
small non-coding RNA-mediated pathways in the
aub
and piwi phenotypes. Recent studies show that this is
indeed the case, and led to the discovery of an entirely
new family of RNAs.
Figure 3 | Role of miR-181 and miR-155 in lymphocyte development. a | Higher
levels of miR-181 in double positive lymphocytes (DP cells) compared with mature
T cells is accompanied by the higher sensitivity of the T-cell receptor to stimulation by
MHC–peptide complexes. As miR
-181 levels decrease during maturation, the activation
threshold of T-cell receptors increases as a result of increased levels of several
phosphatases modulated by miR-181 (see graph).
b | Mir-155-null mice are characterized
by complex defects in homeostasis of the immune system and globally impaired immune
responses. Among the defects that were characterized in detail, the loss of miR-155-
mediated inhibition of the transcription factor c-MAF led to increased production of
interleukin-4 (IL-4) and T helper-2 (Th2) cells. The germinal centre reaction was disrupted,
resulting in impaired T cell-dependent antibody responses (see main text).
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DNA
Genomic
piRNA cluster
piRNA precursor
A
U
Transposable
element
A
A
U
PIWI
AUB
PIWI
AUB
PIWI
AUB
PIWI
AUB
PIWI
AUB
U U U
U U U
5′
AGO3
A
AGO3
A
AGO3
24th–30th
nucleotide
Zucchini
Squash
Zucchini
Squash
U
U
U
AGO3, AUB and PIWI bind a family of thousands
of small non-coding RNAs, called piRNAs
109,110
.
Approximately 80% of the piRNAs in D. melanogaster
are repeat-associated small interfering RNAs (
rasiRNAs),
a family of RNAs with a sequence that corresponds to
or is complementary to transposable elements
110–113
.
piRNAs are single stranded and slightly longer than the
other known small non-coding RNAs (24–29 nucleo-
tides in D. melanogaster), with a phosphorylated 5′ end
and a 2′-O-methyl (2′-O-me) modification at their
3′ ends, similar to siRNAs (but unlike miRNAs)
113–115
.
Flies that lack a functional pimet gene (the homo-
logue of Arabidopsis thaliana HEN1), which encodes
the enzyme responsible for the 2′-O-methylation of
piRNAs, are viable and fertile but show defects in the
ability of piRNAs to repress retrotransposons
115
. In
D. melanogaster
, piRNA sequences are clustered in dis-
crete sites that are located in areas of pericentromeric
and subtelomeric heterochromatin, which are enriched
in repetitive sequences that derive from transposable
elements
111
. Although the biogenesis of piRNAs has not
yet been entirely characterized, it requires Piwi proteins
but, unlike miRNAs and siRNAs, is not affected by
deletion of Dicer
109
(BOX 2). The role of piRNAs in the
repression of transposable genetic elements was demon-
strated by the simultaneous increase of transposons and
disappearance of piRNAs in piwi and aub mutants
109
.
Furthermore, previously characterized master regula-
tors of transposon activity, such as Flamenco, coincide
exactly with piRNA clusters
111
.
Disruption of the rasiRNA pathway also led to defects
in specification of the embryonic axis. Mutations in aub
and two other genes involved in rasiRNA pathways — the
putative helicases Armitage (armi) and Spindle-E (spn‑E)
— resulted in premature expression of the posterior
determinant Oskar
and defects in the polarization of the
microtubule cytoskeleton
116
. However, loss-of-function
mutations of genes encoding ATR and CHK2 kinases,
which function in DNA-damage signalling, suppressed
the embryonic axis defects of armi and aub, but not the
defects in transposon suppression
117,118
. Furthermore,
armi and aub mutants accumulated DNA breaks, as
shown by increased accumulation of foci of the histone
γ-H2Av
117
. These findings indicated that the rasiRNA
pathway does not directly control axis specification in
D. melanogaster
embryos, which was perturbed in armi
and aub mutants as a result of increased DNA damage and
activation of ATR and CHK2, possibly caused by
increased transposon mobilization
117
.
piRNAs in vertebrate germline development. The piRNA
pathway in vertebrates shows similarities to the rasiRNA
pathway in D. melanogaster, as well as some intriguing
differences. Similar to flies, mice have three genes that
belong to the Piwi family, called Mili, Miwi and Miwi2,
which are expressed exclusively in the testis at different
stages of development. The zebrafish piwi orthologue
ziwi is also expressed in the ovaries
119
. In the mouse, Mili
is expressed in mitotic spermatogonia and disappears
towards the end of prophase of the first meiosis, whereas
Miwi appears during prophase of the first meiosis. As
in D. melanogaster, null deletion of these genes caused
profound defects in gametogenesis, albeit only in males.
In Miwi‑ and Mili‑null mutants, spermatogenesis was
arrested during meiosis, with defects appearing earlier
in Mili mutants, consistent with its early expression
120,121
.
Meiotic defects were observed in Miwi2 mutants, along
Box 2 | Generation of piRNAs
The peculiar features of the Piwi-interacting RNAs (piRNAs) that are connected with
individual members of the Piwi family of Argonaute proteins have led to a model for the
generation and amplification of piRNAs (see figure). In Drosophila melanogaster, most of
the piRNAs that co-purify with PIWI (P-element induced wimpy testis) and AUB
(Aubergine) proteins are in the antisense orientation to functional transposons, whereas
the piRNAs that co-purify with AGO3 are mostly in the sense orientation; therefore,
extensive complementarity exists between the AGO3-associated and AUB/PIWI-
interacting pools of piRNAs
111,112
. Interestingly, the 5′ ends of complementary piRNAs
that are associated with AGO3 and AUB/PIWI are separated by ten nucleotides.
Consistent with the complementarity of the first ten nucleotides, most piRNAs that are
associated with AUB and PIWI carry a uridine residue at their 5′ end, whereas there is a
strong bias for adenosine residues at the tenth position in piRNAs from the pool that
co-purifies with AGO3. These features imply that piRNAs are amplified and maintained
by a mechanism that is different from the biogenesis of small interfering RNAs and
microRNAs, which relies on Dicer activity.
According to the proposed model, antisense primary piRNAs, which are generated
from fragments of transposons in the genome, associate with AUB and PIWI proteins
and target complementary active transposons. AUB and PIWI cut the target transposon
at the residue that is complementary to the tenth nucleotide of the piRNA, generating
the 5′ end of a secondary piRNA. Subsequent processing — possibly involving the
putative nucleases Zucchini and Squash that cleave it 24–29 nucleotides downstream
143
— generates secondary piRNAs in the sense orientation, which are bound by AGO3.
The AGO3–piRNA complex can then generate new antisense piRNAs from primary
transcripts that are encoded from the piRNA-generating clusters
111,112
. Thus, sites of
integration of defective transposons (such as the flamenco locus), from which primary
antisense piRNAs are generated, might serve as a genetic memory of previous invasions
by parasitic genetic elements
111
.
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rasiRNA
Repeat-associated small
interfering RNA that is derived
from highly repetitive genomic
loci. rasiRNA is involved in the
establishment and
maintenance of
heterochromatin and
transposon control.
Seminiferous tubules
Structures in the testis where
spermatocytes mature.
with the disappearance of spermatogonia in adult animals,
resulting in completely empty seminiferous tubules
122
.
Similar to D. melanogaster, the estimated 2 × 10
5
potential vertebrate piRNAs are characteristically
longer than miRNAs and siRNAs, and are encoded by a
small number of genomic clusters
119,123–128
. Within each
cluster, all piRNAs are encoded from the same strand,
suggesting the possibility of a primary transcript that
encompasses the cluster, but with no predicted hair-
pin structures, unlike miRNA primary transcripts.
Furthermore, although the genomic location of the clus-
ters is conserved between mammals, the sequences of
the single piRNAs are not. Interestingly, important dif-
ferences exist between the piRNAs that are expressed in
early spermatogenesis (when Mili is expressed) and the
piRNAs that are expressed after the first meiosis
129
. Mili-
associated piRNAs contain many sequences that match
transposable elements and these piRNAs are encoded by
genomic clusters that are rich in nested, often defective,
transposon sequences, similar to the master regulators
of transposon activity in D. melanogaster
129
. Unexpectedly,
only ~17% of the clusters encoding piRNAs that are
expressed late in spermatogenesis are located within
repeated sequences, which is less than the percentage
expected by chance, given that ~40% of the mouse
genome is made up of such sequences
123,124
.
Consistent with a conserved role in controlling
the mobilization of transposable elements, both Mili-
and Miwi2-null mouse mutants show increased levels
of active transposons. In contrast to the situation in
D. melanogaster
, the increased mobilization of trans-
posable elements in Mili and Miwi2 mutants was
accompanied by decreased DNA methylation of the
mobilized elements, suggesting the possibility of a
functional relationship between Piwi and DNA meth-
ylation
122,129
. Furthermore, analysis of early piRNAs
complementary to transposable elements shows an
enrichment of stretches of precisely ten complemen-
tary nucleotides starting from the 5′ end; recipro-
cally, a significant enrichment of adenosine residues
was detected in position 10, matching the uridine
residue at the 5′ end of most piRNAs. These features
are similar to D. melanogaster piRNAs and allow the
hypothesis about the feed-forward loop of transposon
degradation and piRNA amplification to be extended to
vertebrates
129
(BOX 2).
Thus, shared features of piRNAs in D. melanogaster
and vertebrates suggest a common mechanism of action
that involves the control of transposable elements.
Nonetheless, most piRNAs in mouse spermatocytes do
not match transposable element sequences and their
sequences are not conserved. It is therefore plausible
that many piRNAs in the mouse act through entirely
different mechanisms or regulate different biological
functions altogether.
Concluding remarks
The targeted deletion of genes in mice has provided an
invaluable strategy to understand the functional role of
protein-coding genes. Null mutations in invertebrate
miRNA genes resulted, in some cases, in dramatic
developmental phenotypes, while in other cases, only
the simultaneous deletion of more than one function-
ally related miRNA resulted in appreciable pheno-
types
5,19,152
. Given the large number of duplications of
miRNA-encoding sequences in vertebrate genomes,
and the presence of large families of miRNAs that are
similar in their sequence and pattern of expression,
functional redundancy is to be expected to an even
higher degree in mammals. By contrast, some miRNAs
are likely to affect the expression of a large number of
functionally related protein-encoding genes, and their
absence is expected to result in profound phenotypes.
The recent descriptions of mice that carry deletions
of single miRNAs provide great insight into the roles of
these molecules in vivo.
In the future, genetic deletion of single or multiple
miRNA or piRNA loci is likely to become an essential
aspect of the functional analysis of small non-coding
RNAs, as it has been in the past two decades for pro-
tein-coding RNAs. As demonstrated by the example
of miR-208 in mice, or of miR-14 in D. melanogaster,
miRNA deletion can result in increased vulnerability
to stress conditions, which might be difficult to assess
under standard laboratory conditions
14,82
(reviewed
in REF. 130). One specific challenge in assessing the
role of miRNAs that are involved in the maintenance
of homeostasis in the face of external stimuli will be to
devise experimental assays that mimic aspects of the
complexity of life in a natural environment.
Furthermore, the diverse phenotypes that are associ-
ated with genetic deletions of miRNAs generally derive
from increased expression of their target genes. Another
challenge for research in this field in the next few years
will be the reliable identification of the in vivo mRNA
targets of miRNAs. Although computational methods
of miRNA target prediction have rapidly improved,
experimental methods for the reliable identification
of regulated mRNAs could greatly foster our under-
standing of miRNA function in vivo. In addition, these
studies are likely to focus attention on small RNA genes
as important loci in various aspects of human disease,
including birth defects, cardiac arrhythmia, organ
failure and the different forms of neoplasia.
1. Jacob, F. & Monod, J. Genetic regulatory mechanisms
in the synthesis of proteins. J. Mol. Biol. 3, 318–356
(1961).
2. Esquela-Kerscher, A. & Slack, F. J. Oncomirs —
microRNAs with a role in cancer. Nature Rev. Cancer
6, 259–269 (2006).
3. Lee, R. C., Feinbaum, R. L. & Ambros, V. The
C. elegans heterochronic gene lin-4 encodes small
RNAs with antisense complementarity to lin-14.
Cell 75, 843–854 (1993).
4.
Wightman, B., Ha, I. & Ruvkun, G. Posttranscriptional
regulation of the heterochronic gene lin-14 by lin-4
mediates temporal pattern formation in C. elegans.
Cell 75, 855–862 (1993).
5. Reinhart, B. J. et al. The 21-nucleotide let-7 RNA
regulates developmental timing in Caenorhabditis
elegans. Nature 403, 901–906 (2000).
6. Yoo, A. S. & Greenwald, I. LIN-12/Notch activation
leads to microRNA-mediated down-regulation of
Vav in C. elegans. Science 310, 1330–1333 (2005).
7. Johnson, S. M. et al.
RAS is regulated by the
let-7 microRNA family.
Cell 120, 635–647
(2005).
8.
Hobert, O. Architecture of a microRNA-controlled
gene regulatory network that diversifies neuronal cell
fates. Cold Spring Harb. Symp. Quant. Biol. 71,
181–188 (2006).
9. Stark, A., Brennecke, J., Russell, R. B. & Cohen, S. M.
Identification of Drosophila microRNA targets. PLoS
Biol. 1, e60 (2003).
REVIEWS
228
|
MARCH 2008
|
VOLUME 9 www.nature.com/reviews/molcellbio
© 2008 Nature Publishing Group

10. Leaman, D. et al. Antisense-mediated depletion
reveals essential and specific functions of microRNAs
in Drosophila development. Cell 121, 1097–1108
(2005).
11.
Ronshaugen, M., Biemar, F., Piel, J., Levine, M. &
Lai, E. C. The
Drosophila microRNA iab-4 causes a
dominant homeotic transformation of halteres to
wings. Genes Dev. 19, 2947–2952 (2005).
12. Brennecke, J., Hipfner, D. R., Stark, A., Russell, R. B.
& Cohen, S.
M. Bantam encodes a developmentally
regulated microRNA that controls cell proliferation
and regulates the proapoptotic gene hid in Drosophila.
Cell 113, 25–36 (2003).
13. Thompson, B. J. & Cohen, S. M. The Hippo pathway
regulates the bantam microRNA to control cell
proliferation and apoptosis in Drosophila. Cell 126,
767–774 (2006).
14. Xu, P., Vernooy, S. Y., Guo, M. & Hay, B. A. The
Drosophila microRNA Mir-14 suppresses cell death
and is required for normal fat metabolism. Curr. Biol.
13, 790–795 (2003).
15. Lai, E. C., Tam, B. & Rubin, G. M. Pervasive regulation
of Drosophila Notch target genes by GY-box-,
Brd-box-, and K-box-class microRNAs.
Genes Dev. 19,
1067–1080 (2005).
16. Li, X. & Carthew, R. W. A microRNA mediates EGF
receptor signaling and promotes photoreceptor
differentiation in the Drosophila eye. Cell 123,
1267–1277 (2005).
17. Varghese, J. & Cohen, S. M. microRNA miR-14 acts to
modulate a positive autoregulatory loop controlling
steroid hormone signaling in Drosophila. Genes Dev.
21, 2277–2282 (2007).
18. Bartel, D. P. MicroRNAs: genomics, biogenesis,
mechanism, and function. Cell 116, 281–297 (2004).
19. Abbott, A. L. et al. The let-7 microRNA family
members mir
-48, mir-84, and mir-241 function
together to regulate developmental timing in
Caenorhabditis elegans. Dev. Cell 9, 403–414 (2005).
20. Bushati, N. & Cohen, S. M. MicroRNA functions.
Annu. Rev. Cell Dev. Biol. 23, 175–205 (2007).
21. Kloosterman, W. P. & Plasterk, R. H. The diverse
functions of microRNAs in animal development and
disease. Dev. Cell 11, 441–450 (2006).
22. Molnar, A., Schwach, F., Studholme, D. J.,
Thuenemann, E.
C. & Baulcombe, D. C. miRNAs
control gene expression in the single-cell alga
Chlamydomonas reinhardtii. Nature 447,
1126–1129 (2007).
23.
Zhao, T. et al. A complex system of small RNAs in the
unicellular green alga Chlamydomonas reinhardtii.
Genes Dev. 21, 1190–1203 (2007).
24. Lee, Y., Han, J., Yeom, K. H., Jin, H. & Kim, V. N.
Drosha in primary microRNA processing. Cold Spring
Harb. Symp. Quant. Biol. 71, 51–57 (2006).
25. Ruby, J. G., Jan, C. H. & Bartel, D. P. Intronic
microRNA precursors that bypass Drosha processing.
Nature 448, 83–86 (2007).
26. Okamura, K., Hagen, J. W., Duan, H., Tyler, D. M. &
Lai, E.
C. The miRtron pathway generates microRNA-
class regulatory RNAs in Drosophila. Cell 130,
89–100 (2007).
27. Berezikov, E., Chung, W. J., Willis, J., Cuppen, E. &
Lai, E. C. Mammalian mirtron genes.
Mol. Cell 28,
328–336 (2007).
28. Pillai, R. S., Bhattacharyya, S. N. & Filipowicz, W.
Repression of protein synthesis by miRNAs: how
many mechanisms? Trends Cell Biol. 17, 118–126
(2007).
29. Lewis, B. P., Shih, I. H., Jones-Rhoades, M. W., Bartel,
D.
P. & Burge, C. B. Prediction of mammalian
microRNA targets. Cell 115, 787–798 (2003).
30. Rajewsky, N. & Socci, N. D. Computational
identification of microRNA targets. Dev. Biol. 267,
529–535 (2004).
31.
Grimson, A. et al. MicroRNA targeting specificity in
mammals: determinants beyond seed pairing.
Mol. Cell 27, 91–105 (2007).
32.
Long, D. et al. Potent effect of target structure on
microRNA function. Nature Struct. Mol. Biol. 14,
287–294 (2007).
33.
Zhao, Y., Samal, E. & Srivastava, D. Serum response
factor regulates a muscle-specific microRNA that
targets Hand2 during cardiogenesis. Nature 436,
214–220 (2005).
34. Vella, M. C., Reinert, K. & Slack, F. J. Architecture of a
validated microRNA::target interaction. Chem. Biol.
11, 1619–1623 (2004).
35.
Landgraf, P. et al. A mammalian microRNA expression
atlas based on small RNA library sequencing.
Cell 129, 1401–1414 (2007).
36.
Rigoutsos, I. et al. Short blocks from the noncoding
parts of the human genome have instances within
nearly all known genes and relate to biological
processes. Proc. Natl Acad. Sci. USA 103,
6605–6610 (2006).
37.
Berezikov, E. et al. Phylogenetic shadowing and
computational identification of human microRNA
genes. Cell 120, 21–24 (2005).
38.
Xie, X. et al. Systematic discovery of regulatory motifs
in human promoters and 3′ UTRs by comparison of
several mammals. Nature 434, 338–345 (2005).
39. Lim, L. P. et al.
Microarray analysis shows that some
microRNAs downregulate large numbers of target
mRNAs. Nature 433, 769–773 (2005).
40. Stark, A., Brennecke, J., Bushati, N., Russell, R. B. &
Cohen, S.
M. Animal microRNAs confer robustness to
gene expression and have a significant impact on
3′ UTR evolution. Cell 123, 1133–1146 (2005).
41. Farh, K. K. et al.
The widespread impact of
mammalian microRNAs on mRNA repression and
evolution. Science 310, 1817–1821 (2005).
42.
Hornstein, E. & Shomron, N. Canalization of
development by microRNAs. Nature Genet. 38
(Suppl.), S20–S24 (2006).
43. Bartel, D. P. & Chen, C. Z. Micromanagers of gene
expression: the potentially widespread influence of
metazoan microRNAs. Nature Rev. Genet. 5,
396–400 (2004).
44.
Bernstein, E. et al. Dicer is essential for mouse
development. Nature Genet. 35, 215–217 (2003).
45.
Kanellopoulou, C. et al. Dicer-deficient mouse
embryonic stem cells are defective in differentiation and
centromeric silencing. Genes Dev. 19, 489–501 (2005).
46.
Wang, Y., Medvid, R., Melton, C., Jaenisch, R. &
Blelloch, R. DGCR8 is essential for microRNA
biogenesis and silencing of embryonic stem cell self-
renewal. Nature Genet. 39, 380–385 (2007).
47. Giraldez, A. J. et al.
MicroRNAs regulate brain
morphogenesis in zebrafish. Science 308,
833–838 (2005).
48. Murchison, E. P. et al.
Critical roles for Dicer in the
female germline. Genes Dev. 21, 682–693 (2007).
49. Muljo, S. A. et al. Aberrant T cell differentiation in the
absence of Dicer. J. Exp. Med. 202, 261–269 (2005).
50.
Tang, F. et al. Maternal microRNAs are essential for
mouse zygotic development. Genes Dev. 21,
644–648 (2007).
51.
Yi, R. et al. Morphogenesis in skin is governed by
discrete sets of differentially expressed microRNAs.
Nature Genet. 38, 356–362 (2006).
52.
Zhao, Y. et al. Dysregulation of cardiogenesis, cardiac
conduction, and cell cycle in mice lacking miRNA-1-2.
Cell 129, 303–317 (2007).
53. Makeyev, E. V., Zhang, J., Carrasco, M. A. &
Maniatis, T. The microRNA miR-124 promotes
neuronal differentiation by triggering brain-specific
alternative pre-mRNA splicing. Mol. Cell 27,
435–448 (2007).
54. Stern, C. D. Evolution of the mechanisms that
establish the embryonic axes. Curr. Opin. Genet. Dev.
16, 413–418 (2006).
55. Faure, S., Lee, M. A., Keller, T., ten Dijke, P. &
Whitman, M. Endogenous patterns of TGFβ
superfamily signaling during early Xenopus
development. Development 127, 2917–2931 (2000).
56.
Agius, E., Oelgeschlager, M., Wessely, O., Kemp, C. &
De Robertis, E. M. Endodermal Nodal-related signals
and mesoderm induction in Xenopus. Development
127, 1173–1183 (2000).
57.
Martello, G. et al. MicroRNA control of Nodal
signalling. Nature 449, 183–188 (2007).
58. Choi, W. Y., Giraldez, A. J. & Schier, A. F. Target
protectors reveal dampening and balancing of nodal
agonist and antagonist by miR-430.
Science 318,
271–274 (2007).
59.
Lipscombe, D. Neuronal proteins custom designed by
alternative splicing. Curr. Opin. Neurobiol. 15,
358–363 (2005).
60. Ule, J. & Darnell, R. B. RNA binding proteins and the
regulation of neuronal synaptic plasticity. Curr. Opin.
Neurobiol. 16, 102–110 (2006).
61. Kosik, K. S. The neuronal microRNA system.
Nature
Rev. Neurosci. 7, 911–920 (2006).
62.
Lagos-Quintana, M., Rauhut, R., Meyer, J., Borkhardt,
A. & Tuschl, T. New microRNAs from mouse and
human. RNA 9, 175–179 (2003).
63. Sempere, L. F. et al.
Expression profiling of
mammalian microRNAs uncovers a subset of brain-
expressed microRNAs with possible roles in murine
and human neuronal differentiation. Genome Biol. 5,
R13 (2004).
64. Spellman, R. & Smith, C. W. Novel modes of splicing
repression by PTB. Trends Biochem. Sci. 31, 73–76
(2006).
65. Boutz, P. L. et al.
A post-transcriptional regulatory
switch in polypyrimidine tract-binding proteins
reprograms alternative splicing in developing neurons.
Genes Dev. 21, 1636–1652 (2007).
66. Boutz, P. L., Chawla, G., Stoilov, P. & Black, D. L.
MicroRNAs regulate the expression of the alternative
splicing factor nPTB during muscle development.
Genes Dev. 21, 71–84 (2007).
67. Conaco, C., Otto, S., Han, J. J. & Mandel, G.
Reciprocal actions of REST and a microRNA promote
neuronal identity. Proc. Natl Acad. Sci. USA 103,
2422–2427 (2006).
68. Coulson, J. M. Transcriptional regulation: cancer,
neurons and the REST. Curr. Biol. 15, R665–R668
(2005).
69. Visvanathan, J., Lee, S., Lee, B., Lee, J. W. & Lee, S. K.
The microRNA miR
-124 antagonizes the anti-neural
REST/SCP1 pathway during embryonic CNS
development. Genes Dev. 21, 744–749 (2007).
70.
Lagos-Quintana, M. et al. Identification of tissue-
specific microRNAs from mouse. Curr. Biol. 12,
735–739 (2002).
71. Kwon, C., Han, Z., Olson, E. N. & Srivastava, D.
MicroRNA1 influences cardiac differentiation in
Drosophila and regulates Notch signaling. Proc. Natl
Acad. Sci. USA 102, 18986–18991 (2005).
72. Sokol, N. S. & Ambros, V. Mesodermally expressed
Drosophila microRNA-1 is regulated by Twist and is
required in muscles during larval growth. Genes Dev.
19, 2343–2354 (2005).
73. Chen, J. F. et al. The role of microRNA-1 and
microRNA
-133 in skeletal muscle proliferation and
differentiation. Nature Genet. 38, 228–233 (2006).
74.
Yang, B. et al. The muscle-specific microRNA miR-1
regulates cardiac arrhythmogenic potential by
targeting GJA1 and KCNJ2. Nature Med. 13,
486–491 (2007).
75.
Clop, A. et al. A mutation creating a potential
illegitimate microRNA target site in the myostatin
gene affects muscularity in sheep. Nature Genet. 38,
813–818 (2006).
76. Olson, E. N. & Schneider, M. D. Sizing up the heart:
development redux in disease. Genes Dev. 17,
1937–1956 (2003).
77.
van Rooij, E. et al. A signature pattern of stress-
responsive microRNAs that can evoke cardiac
hypertrophy and heart failure. Proc. Natl Acad. Sci.
USA 103, 18255–18260 (2006).
78. McCarthy, J. J. & Esser, K. A. MicroRNA-1 and
microRNA
-133a expression are decreased during
skeletal muscle hypertrophy. J. Appl. Physiol. 102,
306–313 (2007).
79. Sayed, D., Hong, C., Chen, I. Y., Lypowy, J. &
Abdellatif, M. MicroRNAs play an essential role in the
development of cardiac hypertrophy. Circ. Res. 100,
416–424 (2007).
80.
Care, A. et al. MicroRNA-133 controls cardiac
hypertrophy. Nature Med. 13, 613–618 (2007).
81.
Cheng, Y. et al. MicroRNAs are aberrantly expressed
in hypertrophic heart: do they play a role in cardiac
hypertrophy? Am. J. Pathol. 170, 1831–1840
(2007).
82.
van Rooij, E. et al. Control of stress-dependent cardiac
growth and gene expression by a microRNA. Science
316, 575–579 (2007).
Demonstrates that miR‑208 is required for cardiac
hypertrophy.
83. Ito, M. & Roeder, R. G. The TRAP/SMCC/Mediator
complex and thyroid hormone receptor function.
Trends Endocrinol. Metab. 12, 127–134 (2001).
84.
Fatica, A. et al. MicroRNAs and hematopoietic
differentiation. Cold Spring Harb. Symp. Quant. Biol.
71, 205–210 (2006).
85. Dahlberg, J. E. & Lund, E. Micromanagement during
the innate immune response. Sci. STKE 2007, pe25
(2007).
86. Chen, C. Z., Li, L., Lodish, H. F. & Bartel, D. P.
MicroRNAs modulate hematopoietic lineage
differentiation. Science 303, 83–86 (2004).
87.
Monticelli, S. et al. MicroRNA profiling of the murine
hematopoietic system. Genome Biol. 6, R71 (2005).
88. Neilson, J. R., Zheng, G. X., Burge, C. B. & Sharp, P. A.
Dynamic regulation of miRNA expression in ordered
stages of cellular development. Genes Dev. 21,
578–589 (2007).
89. Li, Q. J. et al. miR-181a is an intrinsic modulator of
T
cell sensitivity and selection.
Cell 129, 147–161
(2007).
REVIEWS
NATURE REVIEWS
|
MOLECULAR CELL BIOLOGY VOLUME 9
|
MARCH 2008
|
229
© 2008 Nature Publishing Group

Shows that variations in the levels of miR‑181
during T lymphocyte maturation regulates the
sensitivity to TCR activation.
90. Clurman, B. E. & Hayward, W. S. Multiple proto-
oncogene activations in avian leukosis virus-induced
lymphomas: evidence for stage-specific events.
Mol. Cell. Biol. 9, 2657–2664 (1989).
91. Tam, W., Ben-Yehuda, D. & Hayward, W. S.
bic, a novel
gene activated by proviral insertions in avian leukosis
virus-induced lymphomas, is likely to function through
its noncoding RNA. Mol. Cell. Biol. 17, 1490–1502
(1997).
92.
Metzler, M., Wilda, M., Busch, K., Viehmann, S. &
Borkhardt, A. High expression of precursor
microRNA-155/BIC RNA in children with Burkitt
lymphoma. Genes Chromosomes Cancer 39,
167–169 (2004).
93.
Haasch, D. et al. T cell activation induces a
noncoding RNA transcript sensitive to inhibition by
immunosuppressant drugs and encoded by the
proto-oncogene, BIC. Cell. Immunol. 217, 78–86
(2002).
94. O’Connell, R. M., Taganov, K. D., Boldin, M. P., Cheng,
G. & Baltimore, D. MicroRNA
-155 is induced during
the macrophage inflammatory response. Proc. Natl
Acad. Sci. USA 104, 1604–1609 (2007).
95. Iorio, M. V. et al.
MicroRNA gene expression
deregulation in human breast cancer. Cancer Res. 65,
7065–7070 (2005).
96. Eis, P. S. et al. Accumulation of miR-155 and BIC RNA
in human B
cell lymphomas.
Proc. Natl Acad. Sci. USA
102, 3627–3632 (2005).
97.
van den Berg, A. et al. High expression of B-cell
receptor inducible gene BIC in all subtypes of Hodgkin
lymphoma. Genes Chromosomes Cancer 37, 20–28
(2003).
98.
Kluiver, J. et al. Lack of BIC and microRNA miR-155
expression in primary cases of Burkitt lymphoma.
Genes Chromosomes Cancer 45, 147–153 (2006).
99.
Kluiver, J. et al. BIC and miR-155 are highly
expressed in Hodgkin, primary mediastinal and
diffuse large B cell lymphomas.
J. Pathol. 207,
243–249 (2005).
100.
Yanaihara, N. et al. Unique microRNA molecular
profiles in lung cancer diagnosis and prognosis.
Cancer Cell 9, 189–198 (2006).
101. Thai, T. H. et al.
Regulation of the germinal center
response by microRNA-155.
Science 316, 604–608
(2007).
102.
Rodriguez, A. et al. Requirement of bic/microRNA-155
for normal immune function. Science 316, 608–611
(2007).
This study, together with reference 101, describes
the alterations of immune system functions in
Mir‑155‑null mutants.
103.
Tabara, H. et al. The rde-1 gene, RNA interference,
and transposon silencing in C. elegans. Cell 99,
123–132 (1999).
104. Ketting, R. F., Haverkamp, T. H., van Luenen, H. G. &
Plasterk, R.
H. MUT-7 of
C. elegans, required for
transposon silencing and RNA interference, is a
homolog of Werner syndrome helicase and RNaseD.
Cell 99, 133–141 (1999).
105. Lin, H. & Spradling, A. C. A novel group of
pumilio
mutations affects the asymmetric division of germline
stem cells in the Drosophila ovary. Development 124,
2463–2476 (1997).
106. Cox, D. N. et al.
A novel class of evolutionarily
conserved genes defined by piwi are essential for
stem cell self-renewal. Genes Dev. 12, 3715–3727
(1998).
107.
Schupbach, T. & Wieschaus, E. Female sterile
mutations on the second chromosome of Drosophila
melanogaster. II. Mutations blocking oogenesis or
altering egg morphology. Genetics 129, 1119–1136
(1991).
108. Harris, A. N. & Macdonald, P. M. Aubergine encodes a
Drosophila polar granule component required for pole
cell formation and related to eIF2C. Development
128, 2823–2832 (2001).
109. Vagin, V. V. et al.
A distinct small RNA pathway
silences selfish genetic elements in the germline.
Science 313, 320–324 (2006).
110.
Saito, K. et al. Specific association of Piwi with
rasiRNAs derived from retrotransposon and
heterochromatic regions in the Drosophila genome.
Genes Dev. 20, 2214–2222 (2006).
111.
Brennecke, J. et al. Discrete small RNA-generating loci
as master regulators of transposon activity in
Drosophila. Cell 128, 1089–1103 (2007).
112. Gunawardane, L. S. et al.
A slicer-mediated
mechanism for repeat-associated siRNA 5′ end
formation in Drosophila. Science 315, 1587–1590
(2007).
References 111 and 112 provide evidence for the
current model of piRNA generation in
D. melanogaster.
113. Aravin, A. A. et al.
The small RNA profile during
Drosophila melanogaster development. Dev. Cell 5,
337–350 (2003).
114.
Saito, K. et al. Pimet, the Drosophila homolog of
HEN1, mediates 2′-O-methylation of Piwi-interacting
RNAs at their 3′ ends. Genes Dev. 21, 1603–1608
(2007).
115. Horwich, M. D. et al.
The Drosophila RNA
methyltransferase, DmHen1, modifies germline
piRNAs and single-stranded siRNAs in RISC.
Curr. Biol. 17, 1265–1272 (2007).
116. Cook, H. A., Koppetsch, B. S., Wu, J. & Theurkauf, W. E.
The Drosophila SDE3 homolog armitage is required
for Oskar mRNA silencing and embryonic axis
specification. Cell 116, 817–829 (2004).
117.
Klattenhoff, C. et al. Drosophila rasiRNA pathway
mutations disrupt embryonic axis specification
through activation of an ATR/Chk2 DNA damage
response. Dev. Cell 12, 45–55 (2007).
118.
Chen, Y., Pane, A. & Schupbach, T. Cutoff and
aubergine mutations result in retrotransposon
upregulation and checkpoint activation in Drosophila.
Curr. Biol. 17, 637–642 (2007).
119.
Houwing, S. et al. A role for Piwi and piRNAs in germ
cell maintenance and transposon silencing in
zebrafish. Cell 129, 69–82 (2007).
120.
Deng, W. & Lin, H. miwi, a murine homolog of piwi,
encodes a cytoplasmic protein essential for
spermatogenesis. Dev. Cell 2, 819–830 (2002).
121.
Kuramochi-Miyagawa, S. et al. Mili, a mammalian
member of piwi family gene, is essential for
spermatogenesis. Development 131, 839–849
(2004).
122. Carmell, M. A. et al.
MIWI2 is essential for
spermatogenesis and repression of transposons in the
mouse male germline. Dev. Cell 12, 503–514
(2007).
123.
Aravin, A. et al. A novel class of small RNAs bind to
MILI protein in mouse testes. Nature 442, 203–207
(2006).
124. Girard, A., Sachidanandam, R., Hannon, G. J. &
Carmell, M.
A. A germline-specific class of small RNAs
binds mammalian Piwi proteins. Nature 442,
199–202 (2006).
125. Grivna, S. T., Beyret, E., Wang, Z. & Lin, H. A novel
class of small RNAs in mouse spermatogenic cells.
Genes Dev. 20, 1709–1714 (2006).
126. Lau, N. C. et al.
Characterization of the piRNA complex
from rat testes. Science 313, 363–367 (2006).
127.
Watanabe, T. et al. Identification and characterization
of two novel classes of small RNAs in the mouse
germline: retrotransposon-derived siRNAs in oocytes
and germline small RNAs in testes. Genes Dev. 20,
1732–1743 (2006).
128. Betel, D., Sheridan, R., Marks, D. S. & Sander, C.
Computational analysis of mouse piRNA sequence and
biogenesis. PLoS Comput. Biol. 3, e222 (2007).
129. Aravin, A. A., Sachidanandam, R., Girard, A.,
Fejes-Toth, K. & Hannon, G. J. Developmentally
regulated piRNA clusters implicate MILI in transposon
control. Science 316, 744–747 (2007).
Describes features that are specific to the pools of
piRNAs associated with different Piwi proteins in
the mouse, showing similarities and differences
between vertebrate and invertebrate piRNAs.
130. Leung, A. K. & Sharp, P. A. microRNAs: a safeguard
against turmoil? Cell 130, 581–585 (2007).
131. Pillai, R. S. et al.
Inhibition of translational initiation
by Let-7 microRNA in human cells.
Science 309,
1573–1576 (2005).
132.
Kiriakidou, M. et al. An mRNA m
7
G cap binding-like
motif within human Ago2 represses translation. Cell
129, 1141–1151 (2007).
133.
Mathonnet, G. et al. MicroRNA inhibition of
translation initiation in vitro by targeting the cap-
binding complex eIF4F. Science 317, 1764–1767
(2007).
134. Thermann, R. & Hentze, M. W.
Drosophila miR2
induces pseudo-polysomes and inhibits translation
initiation. Nature 447, 875–878 (2007).
135. Chendrimada, T. P. et al.
MicroRNA silencing through
RISC recruitment of eIF6. Nature 447, 823–828
(2007).
136. Olsen, P. H. & Ambros, V. The lin-4 regulatory RNA
controls developmental timing in Caenorhabditis
elegans by blocking LIN-14 protein synthesis after the
initiation of translation. Dev. Biol. 216, 671–680
(1999).
137. Maroney, P. A., Yu, Y., Fisher, J. & Nilsen, T. W.
Evidence that microRNAs are associated with
translating messenger RNAs in human cells. Nature
Struct. Mol. Biol. 13, 1102–1107 (2006).
138. Petersen, C. P., Bordeleau, M. E., Pelletier, J. &
Sharp, P. A. Short RNAs repress translation after
initiation in mammalian cells. Mol. Cell 21, 533–542
(2006).
139. Nottrott, S., Simard, M. J. & Richter, J. D. Human
let-7a miRNA blocks protein production on actively
translating polyribosomes. Nature Struct. Mol. Biol.
13, 1108–1114 (2006).
140. Valencia-Sanchez, M. A., Liu, J., Hannon, G. J. &
Parker, R. Control of translation and mRNA
degradation by miRNAs and siRNAs. Genes Dev. 20,
515–524 (2006).
141.
Wakiyama, M., Takimoto, K., Ohara, O. &
Yokoyama, S. Let-7 microRNA-mediated mRNA
deadenylation and translational repression in a
mammalian cell-free system. Genes Dev. 21,
1857–1862 (2007).
142.
Behm-Ansmant, I. et al. mRNA degradation by
miRNAs and GW182 requires both CCR4:NOT
deadenylase and DCP1:DCP2 decapping complexes.
Genes Dev. 20, 1885–1898 (2006).
143.
Pane, A., Wehr, K. & Schupbach, T. zucchini and
squash encode two putative nucleases required for
rasiRNA production in the Drosophila germline.
Dev. Cell 12, 851–862 (2007).
144.
Xiao, C. et al. MiR-150 controls B cell differentiation
by targeting the transcription factor c-Myb.
Cell 131,
146–159 (2007).
145. Harfe, B. D., McManus, M. T., Mansfield, J. H.,
Hornstein, E. & Tabin, C.
J. The RNaseIII enzyme Dicer
is required for morphogenesis but not patterning of
the vertebrate limb. Proc. Natl Acad. Sci. USA 102,
10898–10903 (2005).
146. Harris, K. S., Zhang, Z., McManus, M. T., Harfe, B. D.
& Sun, X. Dicer function is essential for lung
epithelium morphogenesis. Proc. Natl Acad. Sci. USA
103, 2208–2213 (2006).
147. O’Rourke, J. R. et al.
Essential role for Dicer during
skeletal muscle development. Dev. Biol. 311,
359–368 (2007).
148. Cobb, B. S. et al. T cell lineage choice and
differentiation in the absence of the RNase III enzyme
Dicer. J. Exp. Med. 201, 1367–1373 (2005).
149.
Schaefer, A. et al. Cerebellar neurodegeneration in the
absence of microRNAs. J. Exp. Med. 204,
1553–1558 (2007).
150.
Liu, J. et al. Argonaute2 is the catalytic engine of
mammalian RNAi. Science 305, 1437–1441 (2004).
151.
Morita, S. et al. One Argonaute family member, Eif2c2
(Ago2), is essential for development and appears not
to be involved in DNA methylation. Genomics 89,
687–696 (2007).
152. Miska, E. A. et al.
Most Caenorhabditis elegans
microRNAs are individually not essential for
development or viability. PLoS Genet. 3, e2 (2007).
Acknowledgements
We thank M. Boehm for critical reading of the manuscript.
G.S. was supported by the Anna Fuller Fund and a Sessel
Postdoctoral Fellowship. F.S. was supported by grants from
the National Institutes of Health, the McDonnell Foundation
and the Ellison Medical Foundation.
DATABASES
Entrez Gene: http://www.ncbi.nlm.nih.gov/entrez/query.
fcgi?db=gene
BIC | HAND2 | KCND2 | Mili | Miwi | Miwi2
FlyBase: http://www.flybase.org
Ago3 | aub | piwi
OMIM: http://www.ncbi.nlm.nih.gov/entrez/query.
fcgi?db=OMIM
Burkitt lymphoma | Hodgkin lymphoma
UniProtKB: http://beta.uniprot.org/uniprot
αMHC | βMHC | DGCR8 | Dicer | Drosha | IRX5 | Nodal | nPTB |
MEF | MYOD | PTB | REST | THRAP1
FURTHER INFORMATION
Frank J. Slack’s homepage: http://www.yale.edu/slack
ALL LINKS ARE ACTIVE IN THE ONLINE PDF
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