Taken together, these results support the mechanistic model: At high temperature, SET-25
pathway activity is reduced, resulting in the
derepression of many loci in the genome. After a
return to low temperature, SET-25 activity is restored, but it takes multiple generations for repression to be completely reestablished. Expression
from SET-25–repressed repeats therefore transmits information about a prior environmental
exposure in this species.
In mammals, repressed repetitive elements can
also escape epigenetic reprogramming (23, 24)
with variation in the expression of both individual repeats (25) and multicopy heterochromatic
transgenes (26) being transmitted between generations. In flies, diet- (6) and stress-induced (5)
changes in heterochromatin can also be transmitted for at least one generation. It is possible,
therefore, that environmentally triggered changes
in heterochromatin may provide a general mechanism for the epigenetic transmission of information between generations. It is interesting to
speculate that the inheritance of environmentally
triggered changes in expression from repressed
chromatin may have been coopted to provide
adaptive benefits to an organism.
REFERENCES AND NOTES
1. J. C. Jimenez-Chillaron et al., Diabetes 58, 460–468
2. J. J. Remy, Curr. Biol. 20, R877–R878 (2010).
3. S. F. Ng et al., Nature 467, 963–966 (2010).
4. B. R. Carone et al., Cell 143, 1084–1096 (2010).
5. K. H. Seong, D. Li, H. Shimizu, R. Nakamura, S. Ishii, Cell 145,
6. A. Öst et al., Cell 159, 1352–1364 (2014).
7. E. J. Radford et al., Science 345, 1255903 (2014).
8. D. Martínez et al., Cell Metab. 19, 941–951 (2014).
9. M. A. Jobson et al., Genetics 201, 201–212 (2015).
10. P. Huypens et al., Nat. Genet. 48, 497–499 (2016).
11. A. Klosin, B. Lehner, Curr. Opin. Genet. Dev. 36, 41–49
12. O. Rechavi et al., Cell 158, 277–287 (2014).
13. D. Schott, I. Yanai, C. P. Hunter, Sci. Rep. 4, 7387
14. J. Z. Ni et al., Epigenetics Chromatin 9, 3 (2016).
15. A. Ashe et al., Cell 150, 88–99 (2012).
16. M. J. Luteijn et al., EMBO J. 31, 3422–3430 (2012).
17. M. Shirayama et al., Cell 150, 65–77 (2012).
18. B. A. Buckley et al., Nature 489, 447–451 (2012).
19. S. G. Gu et al., Nat. Genet. 44, 157–164 (2012).
20. Materials and methods are available as supplementary
21. S. Berry, M. Hartley, T. S. Olsson, C. Dean, M. Howard, eLife 4,
22. B. D. Towbin et al., Cell 150, 934–947 (2012).
23. J. A. Hackett et al., Science 339, 448–452 (2013).
24. W. W. Tang et al., Cell 161, 1453–1467 (2015).
25. H. D. Morgan, H. G. Sutherland, D. I. K. Martin, E. Whitelaw,
Nat. Genet. 23, 314–318 (1999).
26. L. Daxinger, E. Whitelaw, Nat. Rev. Genet. 13, 153–162
This work was supported by a European Research Council
Consolidator grant (616434), the Spanish Ministry of Economy and
Competitiveness (BFU2011-26206 and SEV-2012-0208), the AXA
Research Fund, the Bettencourt Schueller Foundation, Agència de
Gestió d’Ajuts Universitaris i de Recerca (AGAUR), Framework
Programme 7 project 4DCellFate (277899), and the European
Molecular Biology Laboratory–Center for Genomic Regulation
(CRG) Systems Biology Program. A.K. was partially supported by a
la Caixa Fellowship. E.C. and T.V. were supported by the Spanish
Ministry of Economy and Competitiveness (BFU2015-70581) and
by an FI AGAUR Ph.D. fellowship to E.C. RNA-seq experiments were
performed in the CRG Genomics core facility. RNA-seq data sets
are deposited in the National Center for Biotechnology Information
Gene Expression Omnibus under accession GSE83528.
Materials and Methods
Figs. S1 to S15
Tables S1 to S4
25 July 2016; resubmitted 21 December 2016
Accepted 24 March 2017
Control of muscle formation by the
fusogenic micropeptide myomixer
Pengpeng Bi,1,2 Andres Ramirez-Martinez,1,2 Hui Li,1,2 Jessica Cannavino,1,2
John R. McAnally,1,2 John M. Shelton,3 Efrain Sánchez-Ortiz,1,2
Rhonda Bassel-Duby,1,2 Eric N. Olson1,2*
Skeletal muscle formation occurs through fusion of myoblasts to form multinucleated
myofibers. From a genome-wide clustered regularly interspaced short palindromic
repeats (CRISPR) loss-of-function screen for genes required for myoblast fusion and
myogenesis, we discovered an 84–amino acid muscle-specific peptide that we call
Myomixer. Myomixer expression coincides with myoblast differentiation and is essential
for fusion and skeletal muscle formation during embryogenesis. Myomixer localizes
to the plasma membrane, where it promotes myoblast fusion and associates with
Myomaker, a fusogenic membrane protein. Myomixer together with Myomaker can
also induce fibroblast-fibroblast fusion and fibroblast-myoblast fusion. We conclude that
the Myomixer-Myomaker pair controls the critical step in myofiber formation during
Skeletal muscle is the largest tissue in the body, accounting for ~40% of human body mass. The formation of skeletal muscle be- gins with the specification of muscle cell fate by the myogenic transcription factors
Pax7 and MyoD, followed by the expression of a
vast number of genes that establish muscle structure and function (1, 2). A fundamental step in
this process is the fusion of mononucleated myoblasts to form multinucleated myofibers (3–7).
Similarly, in response to injury, myogenic progenitor cells within the adult musculature are
activated and fuse to generate new myofibers
(8–10). Whereas many of the initial steps of myoblast fusion are similar to those of other fusogenic cell types (11), the components and molecular
basis of fusion of specific cell types, such as
myoblasts, have not been fully defined.
To identify new regulators of myogenesis, we
performed a genome-wide clustered regularly
interspaced short palindromic repeats (CRISPR)
loss-of-function screen for genes required for differentiation and fusion of C2C12 myoblasts, a
mouse muscle cell line (fig. S1A). We infected
60 million C2C12 myoblasts with a lentiviral library comprising a pool of 130,209 single-guide
RNAs (sgRNAs) and CRISPR-associated protein
9 (Cas9) for CRISPR gene editing (12). Lentivirus
infection was performed at a multiplicity of infection in order to retain a 460-fold representation of the library. After puromycin selection for
2 days, myoblast cultures were switched to differentiation medium (DM) for 1 week so as to promote
myotube formation. The cultures were subjected
to a brief exposure to low trypsin (0.00625%),
which promoted the detachment of myotubes,
leaving mononucleated myoblasts attached to
the dishes. Subsequent treatment with 0.25%
trypsin allowed release of myoblasts. As a control for separation of the myotube and myoblast
populations, we detected myosin heavy chain
by means of Western blot analysis, which was
highly enriched in the myotube population
We enumerated sgRNA representation in myoblast and myotube populations by means of high-throughput sequencing. Genes targeted by multiple
myoblast-enriched sgRNAs were scored on the
basis of their relative abundance. This analysis
revealed numerous genes that were targeted by
multiple independent CRISPR sgRNAs that were
enriched in myoblasts. Because we sought to identify genes specifically required for myoblast differentiation or fusion, we narrowed down this
list by comparing these genes with transcripts that
are up-regulated during differentiation of C2C12
myoblasts (13), as well as Pax7+ and Twist2+
myogenic progenitors (fig. S1C) (14). Five genes fulfilled
1Department of Molecular Biology, Hamon Center for
Regenerative Science and Medicine, University of Texas
Southwestern Medical Center, Dallas, TX 75390, USA. 2Sen.
Paul D. Wellstone Muscular Dystrophy Cooperative Research
Center, University of Texas Southwestern Medical Center,
Dallas, TX 75390, USA. 3Department of Internal Medicine,
University of Texas Southwestern Medical Center, Dallas,
TX 75390, USA.
*Corresponding author. Email: firstname.lastname@example.org