ing the first 30 days but is required in the second
30-day period when it binds to the FMR1 gene and
leads to gene silencing.
We next asked whether FMR1 gene silencing
can occur after the 48- to 51-day time point of the
differentiation protocol. In these experiments, we
inhibited FMR1 silencing by culturing differentiating FXS hESCs in the presence of 1a for 60 days.
Withdrawal of 1a for 10 days triggered FMR1
silencing (fig. S14), which suggests that sustained
1a treatment is required to maintain FMR1 in the
transcriptionally active state. The small molecule
1a appears to function to prevent silencing, rather
than reverse silencing, as application of 1a to cells
with an already silenced FMR1 promoter did not
reverse silencing (fig. S15).
We next mapped the part of the FMR1 transcript that binds to the FMR1 gene. To do this, we
performed ChIRP using biotinylated primers that
hybridize to different regions of the FMR1 transcript (Fig. 4A). Primers that hybridize adjacent
to the CGG repeat pulled down the FMR1 promoter, whereas primers directed elsewhere along
the FMR1 transcript resulted in markedly reduced
pull down (Fig. 4B). Together with the finding that
1a blocks the binding of FMR1 mRNA to the FMR1
gene, these data suggest that the CGG-repeat region of the FMR1 transcript interacts with the
We next mapped the part of the FMR1 gene
that is bound to the FMR1 mRNA. In our previous ChIRP experiments, we measured the binding of the FMR1 mRNA to a portion of the FMR1
promoter that lies 92 to 196 base pairs (bp) upstream of the CGG-repeat sequence. To more
precisely define the binding site, we measured the
ChIRP signal both upstream and downstream of
the genomic CGG-repeat sequence in differentiating FXS hESCs. Because of the high G/C content
of the repeat sequence, binding to this region cannot be tested. We found that the ChIRP signal was
detectable on both sides of the ~1200-bp genomic
CGG repeat and is markedly reduced at sites away
from the repeat (Fig. 4C). This pattern of binding
is consistent with the genomic CGG-repeat sequence being the binding site for the FMR1 mRNA
(see fig. S16).
Some noncoding RNAs have been shown to
interact with promoters indirectly through a protein intermediate (18). To determine if FMR1
mRNA binds to its promoter through a protein
intermediate, we performed ChIRP experiments,
except we treated the cross-linked lysate with
trypsin to digest any protein intermediates. For the
control TERC noncoding RNA (15), the binding
to its target promoters was abolished after trypsin
treatment (Fig. 4D). However, FMR1 mRNA binding to the FMR1 gene was not affected by trypsin
We next asked if FMR1 mRNA binds to the
FMR1 gene by forming an RNA•DNA hetero-
duplex. To test this, we used ribonuclease H
(RNase H), which selectively degrades RNA hy-
bridized to DNA. Treatment of the cross-linked
DNA fragments with RNase H selectively pre-
vented the pull down of the FMR1 promoter but
did not affect the TERC pull downs (Fig. 4D).
These data indicate that FMR1 mRNA binds the
FMR1 gene through a direct RNA•DNA duplex
and does not require a protein intermediate.
The FMR1 mRNA CGG repeat may bind to
the complementary CCG portion of the DNA that
becomes accessible while it is being transcribed.
Indeed, transcription through G-rich sequences
causes stalling in vitro and in vivo (19, 20). Con-
ceivably, the nascent FMR1 CGG-repeat RNA
interacts with the template strand of the unwound
DNA to form a RNA•DNA duplex that is highly
stabilized by its G/C content. This would require
that the CGG-repeat sequence in the RNA ac-
hieves a sufficient length to reach back and interact
with the DNA and may contribute to the require-
ment for >200 CGG repeats for silencing. In ad-
dition, the length of the resulting RNA•DNA
duplex may need to be of sufficient length to
activate downstream pathways that induce FMR1
The initial step in FMR1 silencing is the binding
of the FMR1 mRNA to the genomic repeat. The
inability of the FMR1 transcript to bind to the
DNA before day 45 may relate to DNA accessibility during transcription. The expression of diverse DNA helicases, which are known to regulate
DNA accessibility (21, 22), is reduced during hESC
differentiation (23). Conceivably, helicase activity
may contribute to the temporal interaction of FMR1
mRNA and DNA. However, the exact mechanisms
underlying the temporal nature of FMR1 silencing remain unknown.
The formation of the FMR1 RNA•DNA duplex
coincides with the initiation of epigenetic silencing in the FMR1 gene. Because this causes a drop
in FMR1 mRNA expression, subsequent maintenance of FMR1 silencing is unlikely to be FMR1
mRNA–dependent. It remains to be determined
which mechanisms maintain epigenetic silencing of FMR1 throughout the patient’s lifetime.
FXS hESCs allow the characterization of the
endogenous FMR1 transcript transcribed from
the endogenous promoter. This is important because RNA-directed gene silencing frequently
occurs in cis with the nascent transcript affecting
a promoter within the gene locus (24). Indeed,
only FMR1 and not other CGG repeats in the
genome are silenced in FXS (25). Thus, rather
than overexpressing CGG-repeat RNAs, which
is complicated by plasmid instability, pharmaco-logic targeting of the endogenous CGG repeat
provides insight into its role in promoter silencing.
Our data demonstrate that an mRNA can
mediate promoter silencing and links trinucleotide repeat expansion to a novel form of RNA-directed promoter silencing. Epigenetic changes
are seen in diverse repeat-expansion diseases
(26, 27). The prevalence of repeat expansion–
associated epigenetic changes raises the possibility that aspects of the mRNA-directed gene
silencing pathway described here may contribute to gene expression alterations in other repeat-expansion diseases as well.
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Acknowledgments: We thank members of the Jaffrey lab
for helpful comments and suggestions, E. Ferretti for
assistance with chromatin immunoprecipitation, Q. Zhan for
help with hESC cultures, S.L. Nolin and C. Dobkin for PCR on
premutation lines. hESC lines are available from N.Z. and
Z.R. (Weill Cornell Medical College hESC lines, nizanin@med.
cornell.edu, firstname.lastname@example.org) and S.R.J. (SI-214
hESC line, email@example.com), subject to approval by
the Embryonic Stem Cell Research Oversight Committee
(ESCRO) of Weill Cornell Medical College. M.D.D. previously
consulted for SMaRT (Small Molecule Approach to Targeting
RNA) Therapeutics. A patent (U.S. Patent and Trademark
Office application no. 61/694, 977) was filed on 30 August
2012 by the Scripps Research Institute governing composition
and use of 1f and 1a. This work was supported by the
Tri-Institutional Stem Cell Initiative (Tri-I SCI) grant
2008-019 (S.R.J., N.Z., and Z.R.), New York Stem Cell
Foundation-Druckenmiller Fellowship (D.C.), Life Sciences
Research Foundation Fellowship and Tri-I SCI postdoctoral
fellowship (M.S.C.), and a FRAXA postdoctoral fellowship
(W.-Y. Y.). Portions of this project not involving non-NIH registry
stem cells were supported by NIH R01 MH80420 (S.R.J.)
and NIH R01 GM079235 (M.D.D.).
Materials and Methods
Figs. S1 to S16
Tables S1 to S3
11 September 2013; accepted 30 January 2014