© 2008 Nature Publishing Group http://www.nature.com/nsmb

NEWS AND VIEWS
anecdotal evidence of the ability of green tea toremedy various amyloid diseases, such as lightchain amyloidosis, have begun to appear24.
It remains unclear whether EGCG canremodel mature amyloid fibers in a similar
way to other small molecules7. However,once again there is reason to be optimistic.
EGCG may take advantage of Le Châtelier’s
principle to disassemble amyloids and prevent
conformational conversion by shifting theequilibrium dramatically toward nonamyloid
conformers. Amyloid fibers seem todynamically exchange monomers from theirends by the spontaneous dissociation andreassociation of monomers over a biologically
relevant timeframe (days)25. Given that EGCG
inhibits seeded polymerization, it is possible,
perhaps even probable, that it might inhibit
the reassociation of dissociated monomers
with fiber ends and drive the equilibrium
toward soluble forms.
One must keep in mind, however, that
EGCG binds to unfolded proteins in a
nonselective fashion8, which may haveunanticipated pleiotropic consequencesin vivo. Indeed, this property might explain
the diverse activities attributed to EGCG14.

EGCG might inhibit the amyloidogenesisof many polypeptides that begin in anatively unfolded state. However, such broad
specificity may be undesirable, because
mounting evidence suggests that amyloids
and prions have also been captured during
evolution for beneficial purposes26. Pmel17
amyloids mediate melanosome biogenesis
and particular CPEB prions might promote
synaptic changes associated with memory26.
Nonetheless, the studies by Ehrnhoefer
et al.8 provide an important foundation tounderstand the interactions between small
molecules and natively unfolded proteins,and may facilitate the design of more potent
and selective compounds with activity against
exclusively deleterious amyloids.
ACKNOWLEDGMENTS
We thank A. Gitler, M. Lemmon, S. Vashist, E. Sweeny
and H. Wang for comments on the manuscript.
1. Fink, A.L. Curr. Opin. Struct. Biol. 15, 35–41 (2005).
2. Nelson, R. & Eisenberg, D. Adv. Protein Chem. 73,
235–282 (2006).
3. Skovronsky, D.M., Lee, V.M.-Y. & Trojanowski, J.Q.
Annu. Rev. Pathol. Mech. Dis. 1, 151–170 (2006).
4. Dobson, C.M. Nature 432, 824–828 (2004).
5. Ehrnhoefer, D.E. et al. Hum. Mol. Genet. 15,
2743–2751 (2006).

6. Masuda, M. et al. Biochemistry 45, 6085–6094 (2006).
7. Wang, H. et al. Proc. Natl. Acad. Sci. USA 105,
7159–7164 (2008).
8. Ehrnhoefer, D.E. et al. Nat. Struct. Mol. Biol. 15,
558–566 (2008).
9. Wells, J.A. & McClendon, C.L. Nature 450, 1001–1009(2007).
10. Krishnan, R. & Lindquist, S.L. Nature 435, 765–772(2005).
11. Petkova, A.T. et al. Science 307, 262–265 (2005).
12. Kayed, R. et al. Science 300, 486–489 (2003).
13. Pardridge, W.M. NeuroRx 2, 3–14 (2005).
14. Yang, C.S., Maliakal, P. & Meng, X. Annu. Rev.
Pharmacol. Toxicol. 42, 25–54 (2002).
15. Hammarstrom, P., Wiseman, R.L., Powers, E.T. &Kelly, J.W. Science 299, 713–716 (2003).
16. Bertoncini, C.W. et al. Proc. Natl. Acad. Sci. USA 102,
1430–1435 (2005).
17. Del Mar, C., Greenbaum, E.A., Mayne, L., Englander, S.W.
& Woods, V.L., Jr. Proc. Natl. Acad. Sci. USA 102,
15477–15482 (2005).
18. Conway, K.A. et al. Proc. Natl. Acad. Sci. USA 97,
571–576 (2000).
19. Bitan, G. et al. Proc. Natl. Acad. Sci. USA 100,
330–335 (2003).
20. Cheon, M. et al. PLoS Comput. Biol. 3, e173 (2007).
21. Behrends, C. et al. Mol. Cell 23, 887–897 (2006).
22. Shorter, J. & Lindquist, S. Mol. Cell 23, 425–438(2006).
23. Evans, C.G., Wisen, S. & Gestwicki, J.E. J. Biol. Chem.
281, 33182–33191 (2006).
24. Mereles, D., Wanker, E.E. & Katus, H.A. Clin. Res.
Cardiol. published online, doi:10.1007/s00392-0080649-6 (3 March 2008).
25. Carulla, N. et al. Nature 436, 554–558 (2005).
26. Shorter, J. & Lindquist, S. Nat. Rev. Genet. 6, 435–450(2005).

Endo-siRNAs: yet another layer ofcomplexity in RNA silencing
Timothy W Nilsen
Organisms possessing RNA-dependent RNA polymerase activity are known to produce endogenous smallinterfering RNAs (esiRNAs). It had been thought that organisms such as flies and mammals lacking this activity
would not produce esiRNAs. However, it has now been shown that a functional esiRNA pathway is present insuch animals; the esiRNAs are derived from a variety of endogenous double-stranded RNA substrates.
The world of small-RNA silencing phenomena
has just become markedly larger with the recent
publication of seven complementary papers (fivein Nature1–5, one in Science6 and one in this issueof Nature Structural & Molecular Biology7) thatcollectively document an additional silencing
pathway in Drosophila melanogaster andmammals. This pathway involves the synthesis
and processing of endogenous double-stranded
RNAs (dsRNAs) to yield functional small

Timothy W. Nilsen is at the Center for RNA
Molecular Biology and Department ofBiochemistry, Case Western Reserve University,
School of Medicine, 10900 Euclid Avenue,
Cleveland, Ohio 44106-4973, USA.
e-mail: twn@case.edu

546interfering RNAs (siRNAs) that serve to silencetransposable elements in both germ cells andsomatic tissues, and some specific mRNAs. It isalso possible that these endo-siRNAs, esiRNAs,
have a role in heterochromatin formation
analogous to the function of some endogenous
siRNAs in plants and fission yeast.
To date, esiRNAs have been detected only inorganisms that possess RNA-dependent RNA
polymerases (RDRPs): plants, Caenorhabditis
elegans and fission yeast. Because thesepolymerases transcribe single-stranded RNA(ssRNA) to make dsRNA and are essential forthe production of esiRNAs, it was thoughtthat organisms that did not have RDRPs
would not use an esiRNA pathway owing to
the lack of endogenous dsRNA. However,
it is well known that there are other sourcesof dsRNAs besides those generated by
RDRPs (Fig. 1). These include long hairpin
structures generated by the transcription of
palindromic sequences and dsRNAs generated
by the annealing of complementary RNAs
synthesized by convergent transcription
units. Indeed, these dsRNAs have now beenshown to be the source of esiRNAs in bothD. melanogaster and mice1–7.
In both organisms, extensive studies in manylaboratories have revealed three distinct RNA
silencing pathways, each using distinct small
RNAs and a distinct set of protein factors. The
Piwi-interacting RNA (piRNA) pathway is
involved in silencing transposons in the germline, and piRNAs are bound to the Piwi classof argonaute proteins; it is not yet clear how
these RNAs and proteins exert their silencing

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NEWS AND VIEWS
effects (reviewed in ref. 8). The microRNA(miRNA) pathway is responsible for the posttranscriptional regulation of many mRNAs
via translational repression or enhancement of
miRNA turnover. In D. melanogaster, miRNAs
function in a ribonucleoprotein complex (RNP)
containing Argonaute-1 (AGO1) and recognize
their mRNA targets via imperfect base-pairing.
The biogenesis of miRNAs is well-understood;
long primary transcripts are processed by the
RNase III–like dsRNA nuclease Drosha incollaboration with the dsRNA binding protein
Pasha (reviewed in ref. 9). The Drosha-processed
pre-miRNA is then exported to the cytoplasm
as a pre-miRNA, where it is further processedby another dsRNase, Dicer-1, in collaboration
with another dsRNA binding protein known
as Loquacious. The processed miRNA is thenloaded into AGO1 (reviewed in ref. 10). The
third RNA silencing pathway in D. melanogasteris the small interfering RNA (siRNA) pathway,
which is responsible for detecting and destroying
exogenous dsRNA and any transcripts derived
from the invading dsRNA. This pathway involves
the direct cleavage of the dsRNA by a distinct
dsRNase, Dicer-2, with a distinct partner, r2d2.
Processed dsRNA is then loaded into a distinct
argonaute protein, AGO2. The siRNA pathway
works via perfect homology between the siRNAand target, resulting in cleavage (slicing) of thetarget by AGO2 (reviewed in ref. 11). It is thispathway in both flies and mammalian cells
that is responsible for targeted knockdowns ofyour favorite gene. Until now, it was not known
whether Dicer-2 and AGO2 were simply lying in
wait for an invading dsRNA or had a more activerole in organismal physiology.
To address this question, Czech et al.2 andKawamura et al.3 used immunoprecipitation of
AGO2 in the hope of identifying endogenous
small RNAs associated with it. Both groups
obtained strikingly similar results. Each
identified populations of small RNAs that areclearly distinct from previously characterizedpiRNAs or miRNAs, and both found that mostof these small RNAs, dubbed esiRNAs, were
derived from mobile genetic elements. Other
esiRNAs were processed from either hairpin
structures or from overlapping RNAs formedby convergent transcription.
Three other papers addressed the samequestion via three different routes. Okamura
et al.1 used an informatics approach to predict
long hairpin structures that might give riseto esiRNAs. They were able to identify manyloci that could encode such RNAs, and they
were then able to verify the expression ofsmall RNAs (which they called hpRNAs) thatwere processed from the long hairpins. These
hpRNAs comprise a subset of the esiRNAs
described in Czech et al.2 and Kawamura

et al3. Instead of looking for inverted repeats,
Okamura et al.7 focused on bidirectionaltranscription units in D. melanogaster.
Intriguingly, they found several such loci, andsome but not all yielded esiRNAs. The reasonsfor the selectivity in processing are not clear.
Finally, Ghidiyla et al.6 used yet a different
strategy; they reasoned that any endogenous
siRNAs were likely to be modified at their
3′ ends, as this is where siRNAs produced fromexogenous dsRNA are methylated. Accordingly,
they prepared libraries of small RNAs enrichedby virtue of their 3′ end modifications. They
were thereby able to identify a population ofRNAs that did not correspond to miRNAs or
piRNAs. Similarly to the results reported by
Czech et al.2 and Kawamura et al.3, most of theRNAs were derived from transposable elements;
others clearly were derived from protein-coding
sequences. One unique aspect of this approach
was the discovery of what may be a fourthclass of small RNA, piRNA-like RNAs. Further
experiments will be necessary to elucidate therole(s) of these mysterious RNAs.
Collectively, these five papers provide
compelling evidence of the existence ofendogenous siRNAs in D. melanogaster. Two
additional studies document similar findings inmouse oocytes. Both Tam et al.4 and Watanabe
et al.5 used deep sequencing to examine the small
RNA population in oocytes, and both uncovered,in addition to piRNAs and miRNAs, a family ofwhat seemed to be esiRNAs. These RNAs wereclearly distinct in size from piRNAs and inmany cases were derived from distinct genomic
loci. In this regard, mapping of the esiRNAs
revealed that, much like in D. melanogaster,
the small RNAs were derived from a variety ofsources: long hairpin structures, overlappingtranscription units and transposable elements.
Most intriguingly, a fraction of esiRNAs wereapparently processed from overlapping regionsof functional genes and cognate pseudogenes.
The presence of esiRNAs spanning exon-exonboundaries indicated that spliced mRNAs arecapable of annealing in trans with transcribed
pseudogenes. This exciting finding suggested
that pseudogenes, previously thought to be
nonfunctional, may actually regulate theexpression of their founder gene.
Although the evidence for the existenceof esiRNAs in D. melanogaster and mice(and presumably every other animal) is
overwhelming, do these RNAs have biological
function? All seven papers address this question,and the answer is unambiguously yes. Most
strikingly, esiRNAs have a role (analogous tothat of piRNAs) in suppressing the expressionof mobile genetic elements. Mice deficient for
Dicer (mammals, unlike D. melanogaster, have
only one Dicer gene) show elevated expression

NATURE STRUCTURAL & MOLECULAR BIOLOGY VOLUME 15 NUMBER 6 JUNE 2008

Long hairpins,transcription of
inverted repeats

5′

3′

5′
5′
5′

5′

Convergent
transcription

5′

Symmetricaltranscription

mRNA

Trans-interactions

Transcriptfrom pseudogene
Transposon 1

5′

Transposon 2

Read-throughtranscription oftransposons

Read-through transcription

dsRNA

Figure 1 Sources of endogenous double-stranded
RNAs. See text for details.

of transposable elements. Interestingly, there isa level of specificity in this phenomenon; thatis, only certain transposons seem to be affectedby the esiRNA pathway4,6.
In D. melanogaster, flies deficient in AGO2 also
show increased expression of transposons, andin this organism esiRNAs may be the primarymechanism for silencing mobile elements insomatic cells, which lack the piRNA pathway(reviewed in ref. 8). In addition to their function
in controlling the expression of transposons andother selfish genetic elements, it seems clearthat esiRNAs also regulate specific proteincoding transcripts. Nevertheless, the extent andbiological relevance of this type of regulation
awaits further investigation. In this regard, it
seems possible that the esiRNA pathway may,at least in part, serve a ‘passive’ function in
mammals; that is, the destruction of long cellular
dsRNAs would prevent the inappropriate
activation of cellular defense mechanisms suchas the dsRNA-activated protein kinase (PKR).
Whereas the discovery of esiRNAs is in itself
highly significant, the mechanism by whichthese RNAs are produced is also extremelyinteresting. As noted above, D. melanogaster has
two distinct Dicers, each of which has a ‘partner’dsRNA binding protein: Dicer-2 and r2d2; andDicer-1 and Loquacious. Not surprisingly,
Dicer-2 is responsible for production of
esiRNAs. Remarkably, however, flies deficient in
its partner, r2d2, show no defect in the amountof esiRNAs; rather, mutants in Loquacious
are highly depleted in esiRNAs. This striking547

© 2008 Nature Publishing Group http://www.nature.com/nsmb

NEWS AND VIEWS
observation raises several intriguing questions.
First, how does Dicer-2 ‘know’ which partner
to choose when confronting exogenous andendogenous dsRNA. One possibility is that
there is a subpopulation of Dicer-2, perhaps
post-translationally modified, that specificallyrecruits Loquacious and is devoted to theesiRNA pathway. Further biochemical analysis
may reveal whether this conjecture is correct.
A second mysterious question raised by
the esiRNA pathway is, where in the cell does
Dicer processing take place? In this regard,
Dicers have been thought to be cytoplasmic
enzymes. However, it seems highly unlikelythat the substrates for esiRNA production
would be exported to the cytoplasm forprocessing. Surprisingly, none of the sevenpapers comments on this issue. In the absenceof evidence to the contrary, it seems possible
and perhaps likely that there may be a dedicatednuclear RNAi apparatus in animal cells. If thisproves to be true, it raises the prospect that
small RNAs could be involved in every level ofgene expression in animals; perhaps we are notso different from plants after all.
1. Okamura, K. et al. Nature advance online publication,
doi:10.1038/nature07015 (7 May 2008).
2. Czech, B. et al. Nature advance online publication,

doi:10.1038/nature07007 (7 May 2008).
3. Kawamura, Y. et al. Nature advance online publication,
doi:10.1038/nature06938 (7 May 2008).
4. Tam, O.H. et al. Nature advance online publication,
doi:10.1038/nature06904 (10 April 2008).
5. Watanabe, T. et al. Nature advance online publication,
doi:10.1038/nature06908 (10 April 2008).
6. Ghildiyal, M. et al. Science published online,
doi:10.1126/science.1157396 (10 April 2008).
7. Okamura, K., Balla, S., Martin, R., Liu, N. & Lai, E.C.
Nat. Struct. Mol. Biol. 15, 581–590 (2008).
8. Aravin, A.A., Hannon, G.J. & Brennecke, J. Science
318, 761–764 (2007).
9. Kim, V.N. Nat. Rev. Mol. Cell Biol. 6, 376–385(2005).
10. Zamore, P.D. & Haley, B. Science 309, 1519–15124(2005).
11. Eulalio, A., Huntzinger, E. & Izaurralde, E. Cell 132,
9–11 (2008).

Damage control
Efficient repair of DNA damage is essential to all organisms.
Cells must sense the damage and activate the appropriate
signals to allow DNA to be repaired. If DNA damage is
extensive, the cell might transiently delay the cell cycle toensure the damage is repaired before the cell divides. In more
extreme cases, cell cycle arrest may be permanent, resulting insenescence, or the cell may undergo cell death.
The DNA damage response (DDR) in mammalian cells
comprises early damage sensors, such as the MRN complex(MRE11/Rad50/NBS1), that
detects double-stranded
breaks; mediator proteins,
MDC1 and 53BP1, which
are recruited by MRN; andactivated kinases such as
ATM that phosphorylate thehistone variant H2AX. H2AX
phosphorylation marks thechromatin at the site of DNA
damage and serves as a signal to help recruit the DNA repair
machinery in a complex cascade of events whose sequenceand interdependencies have been difficult to define. Recently,
two groups independently investigated the consequences ofaccumulating specific DDR proteins at a defined DNA site, andfound that the DDR can be activated without DNA damage.
DDR proteins accumulate in distinct DNA repair foci at thesites of DNA lesions. To reconstruct DDR foci in the absenceof DNA damage, both groups fused genes for DDR proteins tothose encoding a fluorescent protein and the bacterial LacR
repressor protein. The fusion proteins were then expressedin cells containing an array of repeats of the LacR target
sequence, lacO, where they became immobilized to this DNA
site, forming a focus that can be visualized by fluorescencemicroscopy. Adding IPTG causes the focus to disassemble, so
one can tell the effects of overproduction of the protein fromthose due to its concentration on a chromatin region.
Soutoglou and Misteli (Science, published online 15 May
2008; doi 10.1126/ science.1159051) found that tetheringof NBS1 led to recruitment of MRE11 as well as MDC1 and
53BP1. Likewise, tethering of MRE11 led to accumulationof NBS1, MDC1 and 53BP1. Moreover, targeting of NBS1,

548

MRE11, MDC1 or ATM was sufficient to activate the DDR inmouse cells, as indicated by several phosphorylation marker
events and cell cycle delay (left hand panel immobolization ofa simple repair factor on chromatin (red), middle panel DDR
activation is indicated by phosphorylation of H2AX (green),right hand panel colocalization). The fact that the DDR canbe triggered without a DNA lesion indicates that although thelesion itself is important for triggering the response, it is not
instrumental for subsequent steps of the DDR. The authors also
provided some insight into who
recruits whom—as expected,
they found that tethered
upstream factors recruited
downstream components,
but they also noticed that
downstream factors suchas MDC1 could recruit
upstream factors, pointing toa feedback loop that might beimportant for amplification of the initial DDR signal. H2AX wasrequired for the checkpoint response, indicating that chromatin
modifications are important for full DDR activation.
Toczyski and colleagues (Mol. Cell 30, 267–276) tethered adifferent set of DDR proteins to yeast chromatin, those recruited
to single-stranded DNA overhangs formed during processing of
double-stranded breaks: the damage-specific DNA clamp, called
the 9-1-1 complex, and the complex Mec1–Ddc2. Mec1 is theyeast ortholog of the mammalian ATR. It phosphorylates theadaptor molecule Rad9 (an ortholog of 53BP1) at the site of DNA
damage, leading to recruitment and activation of the checkpoint
kinase Rad53 (a Chk2 ortholog). Tethering of both the 9-1-1 andthe Mec1-Ddc2 complexes to a chromatin site could activate
the checkpoint responses, with Rad53 phosphorylation anddownstream events including cell cycle arrest occurring in theabsence of DNA damage. Specific chromatin modifications wererequired for efficient checkpoint signaling.
Both these papers used an elegant and powerful technique
to dissect the events involved in DNA damage signaling andcheckpoint responses, and this approach may be useful in studying
other protein complexes involved in DNA-dependent processes.
Inês Chen

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