ROLES OF CUG-BP, ELAV-LIKE FAMILY MEMBER 1 (CELF1), AN RNA BINDING
PROTEIN, DURING VERTEBRATE HEART DEVELOPMENT
by
YOTAM N. BLECH-HERMONI
Submitted in partial fulfillment of the requirements for the degree of Doctor of
Philosophy
Advisor: Dr. Andrea N. Ladd
Program in Cell Biology,
Department of Molecular Biology and Microbiology,
School of Medicine
CASE WESTERN RESERVE UNIVERSITY
January 2015 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the dissertation of
Yotam N. Blech-Hermoni
Candidate for the degree of Doctor of Philosophy*.
Committee Chair
Dr. Donna M. Driscoll
Committee Member
Dr. Alan M. Tartakoff
Committee Member
Dr. Hua Lou
Date of Defense
November 20, 2014
*Published material is included in accordance with contracts and stipulations set
forth by the publishers.
i Table of Contents
List of Tables ...... vii
List of Figures ...... viii
Acknowledgements ...... x
List of Abbreviations ...... xi
Abstract ...... xiv
Chapter 1: Introduction ...... 1
1.1. Post-transcriptional regulation of gene expression ...... 1
1.2. The CUG-BP, Elav-like family (CELF) of proteins ...... 2
1.2.1. Nomenclature ...... 3
1.2.2. Structure and RNA binding ...... 6
1.2.3. The ‘Divergent Domain’ ...... 9
1.2.4. Expression patterns and roles ...... 9
1.3. CELF1 ...... 10
1.3.1. Roles in the cytoplasm ...... 11
1.3.2. Roles in the nucleus ...... 13
1.3.3. Biological functions of CELF1 ...... 15
1.3.4. Mouse models for the study of CELF1 ...... 17
1.4. Model organisms ...... 17
1.4.1. The mouse ...... 18
1.4.2. The chicken ...... 20
1.4.3. The frog ...... 21
1.4.4. Comparative staging ...... 22
ii 1.5. The current work ...... 22
Chapter 2: Materials and Methods ...... 24
2.1. Animal use ...... 24
2.2. Chickens ...... 24
2.3. Chicken embryonic tissue and embryo collection ...... 24
2.4. Chicken embryonic primary cardiomyocyte isolation and culture ...... 25
2.5. Transfection of primary embryonic cardiomyocytes with siRNAs ...... 26
2.6. EdU staining of cultured primary cardiomyocytes ...... 27
2.7. Mice ...... 28
2.8. Mouse embryo collection ...... 29
2.9. Western blotting and nuclear-cytoplasmic fractionation of tissues ...... 29
2.10. Immunofluorescence [IF] ...... 31
2.11. In situ hybridization ...... 32
2.12. Semi-quantitative RT-PCR ...... 33
2.13. Real-time RT-PCR ...... 33
2.14. Cross-linking immunoprecipitation (CLIP) ...... 36
2.15. Analysis of CLIP tags ...... 38
2.16. Frog embryo production ...... 39
2.17. Frog embryo morpholino oligonucleotide (MO) microinjection ...... 40
2.18. Whole-mount immunohistochemistry [IHC] and IF ...... 41
2.19. Bright-field imaging ...... 42
2.20. Frog construct design ...... 42
2.21. Alternative Splicing Assay ...... 43
iii Chapter 3: RNA binding proteins in the regulation of heart development ...... 45
3.1. Introduction ...... 45
3.2. Cardiac cell fate, heart tube formation, and differentiation ...... 48
3.2.1. Formation of the heart tube ...... 53
3.2.2. RBPs in cardiomyocyte differentiation and myofibril development ...... 56
3.3. Cardiac morphogenesis ...... 59
3.3.1. Endocardial cushion development ...... 60
3.3.2. Myocardial trabeculation and compaction ...... 63
3.4. Postnatal maturation of the heart ...... 64
3.4.1. CELF- and MBNL-mediated alternative splicing programs ...... 65
3.4.2. Roles of the SR protein family in the maturing heart ...... 67
3.5. Developmental dysregulation and disease ...... 69
3.5.1. RNA binding proteins and congenital heart defects ...... 70
3.5.2. The recapitulation of fetal programs during adult heart disease ...... 71
3.6. Concluding remarks ...... 73
3.7. Acknowledgements ...... 76
Chapter 4: Diversity and conservation of CELF1 and CELF2 RNA and protein expression patterns during embryonic development ...... 77
4.1. Introduction ...... 77
4.2. Results and Discussion ...... 80
4.2.1. CELF1 and CELF2 undergo dynamic post-transcriptional regulation
during early heart development in chicken ...... 80
4.2.2. Different CELF1 and CELF2 isoforms are expressed in different tissues . 88
iv 4.2.3. Regional distribution of CELF1 and CELF2 in the chicken embryo differ
between tissues ...... 89
4.2.4. The distribution of CELF1 and CELF2 transcripts and CELF1 protein are
largely conserved between chicken and mouse ...... 97
4.3. Concluding remarks ...... 102
4.4. Acknowledgements ...... 104
Chapter 5: CUG-BP, Elav-like family member 1 (CELF1) regulates MYH7B
expression in cardiac muscle ...... 105
5.1. Introduction ...... 105
5.2. Results ...... 108
5.2.1. CELF1 binds predominantly to UG-rich intronic sequences in the
embryonic heart ...... 108
5.2.2. Alternative splicing of MYH7B inversely correlates with total transcript
levels and CELF1 expression ...... 117
5.2.3. CELF-mediated alternative splicing of Mef2a correlates with Myh7b levels
in transgenic mice ...... 122
5.3. Discussion ...... 123
5.4. Concluding remarks ...... 128
5.5. Acknowledgements ...... 130
Chapter 6: CUG-BP, Elav-like family member 1 (CELF1) regulates multiple
aspects of myocardial development ...... 131
6.1. Introduction ...... 131
6.2. Results ...... 133
v 6.2.1. CELF1 is expressed in cardiomyocytes and is predominantly nuclear in
cardiac tissue in vivo ...... 133
6.2.2. Cultured chicken primary embryonic cardiomyocytes recapitulate
expression of cardiac markers and CELF1 seen in the developing heart ...... 136
6.2.3. Knockdown of CELF1 in cultured primary embryonic cardiomyocytes
leads to myofibrillar disorganization ...... 141
6.2.4. CELF1 is required for early events in myofibril assembly in cultured cells
...... 144
6.2.5. CELF1 expression patterns are conserved between frog and chicken .... 152
6.2.6. Knockdown of Celf1 in frog embryos results in cardiac looping defects
and cardiac dysmorphia ...... 155
6.3. Discussion ...... 162
6.4. Acknowledgements ...... 165
Chapter 7: Concluding remarks ...... 167
7.1. Expression patterns and activities of CELF proteins ...... 167
7.2. CELF1 in myofibrillar organization and cardiac morphogenesis ...... 169
7.3. CELF1 in cardiac function of the embryonic heart ...... 172
7.4. Future directions ...... 173
Appendix ...... 177
References ...... 194
vi List of Tables
Table 1. Names previously used for members of the CELF protein family in different
species...... 4
Table 2. Comparative timeline of cardiac morphogenesis in mouse, chicken, and frog...... 19
Table 3. Antibodies used in this work ...... 30
Table 4. Primers and Probes ...... 34
Table 5. Summary of CELF1 and CELF2 expression in embryonic and adult tissuesa
...... 81
Table 6. Lengths of introns containing CLIP tags and relative position of tags within introns ...... 116
Table S1. CELF1 CLIP tags obtained from embryonic day 8 chicken heart...... 177
Table S2. The subset of intronic CELF1 CLIP tags that fall within annotated genes.
...... 177
vii List of Figures
Figure 1. A dendrogram showing the evolutionary relationship between members of
the CELF proteins family...... 7
Figure 2. Mechanisms of RBP-mediated post-transcriptional regulation...... 46
Figure 3. RNA binding proteins have been implicated in the formation, morphogenesis, and maturation of the heart...... 49
Figure 4. Domain structure of RNA binding proteins implicated in regulation of heart development...... 51
Figure 5. CELF1 and CELF2 expression in chicken heart during embryonic development...... 83
Figure 6. Expression of CELF1 and CELF2 proteins in a panel of chicken embryonic tissues...... 87
Figure 7. CELF1 and CELF2 in situ hybridization in chicken embryos...... 90
Figure 8. CELF1 and CELF2 are differentially expressed in the embryonic chicken eye...... 93
Figure 9. Tissue distribution of CELF1 protein in embryonic chicken...... 95
Figure 10. Celf1 and Celf2 in situ hybridization in sagittal sections of mouse embryos.
...... 98
Figure 11. Tissue distribution of CELF1 protein in embryonic mouse...... 101
Figure 12. Cross-linking immunoprecipitation (CLIP) of CELF1 from embryonic chicken heart...... 109
Figure 13. CELF1 CLIP tags are enriched with UG motifs...... 111
Figure 14. Sequence enrichment in Nova CLIP tags differs from CELF1 CLIP tags. 114
viii Figure 15. CELF1 regulates the inclusion of an unannotated exon in chicken MYH7B
transcripts...... 119
Figure 16. Model for direct and indirect regulation of MYH7B expression via CELF- mediated alternative splicing...... 129
Figure 17. Isolated chicken primary embryonic cardiomyocytes represent a highly enriched cardiomyocyte culture system...... 135
Figure 18. Expression of CELF1 over days in culture and development of myofibrillar structure...... 139
Figure 19. Transfection of cultured chicken primary embryonic cardiomyocytes with siRNAs results in robust knockdown of CELF1 protein and transcript levels...... 142
Figure 20. Knockdown of CELF1 using siRNAs leads to reduced cell proliferation and profound disorganization of myofibrillar structure in cultured chicken primary embryonic cardiomyocytes...... 143
Figure 21. Following knockdown of CELF1 at 24 hpp, myofibrillar disorganization persists...... 147
Figure 22. Developmental expression and activity of Celf1 are conserved in Xenopus laevis...... 151
Figure 23. Morpholino-mediated knockdown of celf1 in Xenopus laevis leads to cardiac looping defects, ventral edema, and gut malformation...... 157
Figure 24. Morpholino-mediated knockdown of Celf1 in Xenopus laevis leads to myofibril disorganization in the developing heart...... 161
ix Acknowledgements
I would like to extend my sincere gratitude to the members of my Committee
– Dr. Donna M. Driscoll, Dr. Alan M. Tartakoff, and Dr. Hua Lou – for their dedication
and commitment to my progress and success over the past very many years. I chose
them with confidence that they will rigorously review my work and my progress,
will work well with each other, and will have my short-term and long-term success
in mind. They did not disappoint.
I would like to thank my Advisor, Dr. Andrea N. Ladd, for her unfaltering
professionalism, mentorship, and cheer. I can only hope to one day be for someone
else the kind of mentor she has been to me. Beyond her vision and motivation, I
have benefited greatly from Andrea’s (often-uncontrollable) meticulousness. The
effects of this quality can be seen in my work at the bench as well as in my record
keeping, which would both look quite different if left to my own devices. However,
this quality also extends to her editing skills. Good editors are hard to find and I
have been lucky to work for one.
I would be remiss if I did not thank my friends of the past five years for their
camaraderie and support. It saddens me to leave them, as I have done my best to
avoid this outcome. In no particular order, I thank Tat To Trinh (the beautiful and
adorable), Kien Nguyen, Liz Liu, and Gaurav Choudhary. In fact, and at his request,
this page is dedicated to Gaurav Choudhary.
My deepest gratitude goes out to Dr. Khadijah A. Mitchell, whose example overwhelmingly inspired my participation in leadership and service activities. The
CWRU School of Medicine and Graduate Student Senate owe her a debt of gratitude.
x List of Abbreviations
ACTA1, skeletal actin alpha 1
ACTN2, sarcomeric alpha-actinin
AVC, atrioventricular canal
CELF, CUG-BP, Elav-like family
CELF1, CUG-BP, Elav-like family member 1
CELF2, CUG-BP, Elav-like family member 2
CHAMP, cardiac helicase activated by MEF2 protein;
CLIP, cross-linking immunoprecipitation
Csm, cardiac-specific isoform of Mov10l1
DGCR8, DiGeorge Syndrome critical region gene 8
DGS, DiGeorge Syndrome
DM, dystrophia myotonica (myotonic dystrophy)
DM1, myotonic dystrophy type 1
EdU, 5-ethynyl-2’-deoxyuridine
EMT, epithelial-to-mesenchymal transition
ESRP, epithelial splicing regulatory protein
FHL2, four and a half LIM domains 2
FXR1, Fragile X mental retardation autosomal homolog 1
GAPDH, Glyceraldehyde-3-phosphate dehydrogenase
HERMES, heart and RRM expressed sequence hnRNP, heterogeneous nuclear ribonucleoproteins how, held out wings
xi KH domain, hnRNP K homology domain
MBNL, muscleblind-like
MEF2, myocyte enhancer factor 2
MET, mesenchymal-to-epithelial transition miRNA, microRNA
MYH7B, myosin heavy chain 7B
MYOM, myomesin
NMD, nonsense mediated decay
OCT, optical coherence tomography
OFT, outflow tract
PCR, polymerase chain reaction
PTB, polypyrimidine tract binding protein qPCR, quantitative PCR (real-time PCR)
RBFOX, RNA binding Fox-1 homolog
RBM, RNA binding motif
RBP, RNA binding protein
RISC, RNA-induced silencing complex
RRM, RNA recognition motif
RNA, ribonucleic acid
RS domain, arginine/serine-rich domain
RT-PCR, reverse-transcriptase PCR siRNA, small interfering RNA
SRF, serum response factor
xii SRSF, serine/arginine-rich splicing factor
STAR, signal transduction and activation of RNA
TNNT2, cardiac troponin T
TPM2, Tropomyosin
TTN, titin
UCSC, University of California, Santa Cruz
UTR, untranslated region
xiii Roles of CUG-BP, Elav-Like Family Member 1 (CELF1), an RNA Binding Protein,
During Vertebrate Heart Development
Abstract
by
YOTAM BLECH-HERMONI
The heart is the first organ to become fully functional during embryonic
development. During tissue development, proteins undergo dynamic ebbs and
flows, as genes are induced and repressed. Although RNA binding proteins (RBPs)
intimately interact with target transcripts and mediate their processing, their
involvement in cardiac development has been largely underappreciated. CUG-BP,
Elav-like family member 1 (CELF1) is a multi-functional RBP found in various
tissues, including the myocardium of the developing heart, and has been implicated
in both nuclear and cytoplasmic roles. Currently, little is known about the roles of
CELF1 in heart tissue and less is known about its involvement in normal
organogenesis. In order to address the roles of CELF1 during embryonic
development, I first investigated the expression patterns of two closely related CELF proteins (CELF1 and CELF2) in mouse and chicken embryos, demonstrating the diversity of isoforms and expression patterns of these transcripts and proteins, and the evolutionary conservation of these patterns during vertebrate embryonic development. Targets of CELF1 regulation during heart development are not known.
xiv I report here the identification of MYH7B (a sarcomeric myosin heavy chain) as a novel direct target of CELF1 regulation in the embryonic chicken heart. The regulation of MYH7B by CELF1 demonstrates the versatility and regulatory complexity of RBPs. Based on these findings, as well as published data implicating
CELF1 in cardiac contraction and contractility post-natally, I hypothesized that
CELF1 is involved in the structure and function of the myocardial contractile apparatus in the embryonic heart. I describe investigations of roles of CELF1 during cardiac morphogenesis at the cell, tissue, and embryo levels. I report that CELF1 is important for myofibrillar organization in cultured primary chicken embryonic cardiomyocytes (the only cells in the developing heart in which CELF1 is expressed), as well as for the maintenance of proliferative capacity in these cells. Finally, I evaluate the roles of Celf1 in the developing heart by knocking down the protein in the frog embryo, demonstrating that Celf1 has a role in cardiac morphogenesis (e.g., cardiac looping) and in the general morphology of the heart.
xv Chapter 1: Introduction
1.1. Post-transcriptional regulation of gene expression
The ‘Central Dogma’ posits that deoxyribonucleic acid (DNA; the genome) is transcribed into ribonucleic acid (RNA; the transcriptome), which is then translated into protein (the proteome). The expression of genes into proteins is regulated at every step of this process. Modifications to DNA (e.g., methylation) and chromatin
(e.g., histone modifications), as well as control of levels and availability of transcription factors and their binding, are used to dynamically regulate the levels of transcription. However, decisions of which proteins will be translated and where in the cell they will be produced are largely regulated after the precursor mRNA
(pre-mRNA) is transcribed — i.e., post-transcriptionally. RNA binding proteins
(RBPs) are in contact with transcripts from the time they are transcribed to the moment they are degraded and regulate their identities and availabilities in diverse ways. As a category, post-transcriptional regulation underlies much of the increased complexity seen in the eukaryotic proteome, when compared to the relatively modest change in the number of genes throughout evolution. By far the largest effect on proteome diversification in higher eukaryotes is imparted by RBP- mediated regulation of pre-mRNA alternative splicing (Maniatis and Tasic, 2002).
This process refers to the choice of which sequences are included in the mature mRNA and which are excised, and is estimated to affect over 95% of multi-exon genes in humans (Pan et al., 2008; Wang et al., 2008). This mechanism leads to alternative protein-coding sequences resulting in altered protein function or
1 localization, frameshifts resulting in transcript degradation, or alternative UTRs that
alter translation regulation. A recent focus of intense research has been the
regulation of gene expression by small RNAs (such as micro-RNAs); a process
mediated by RBPs (for example, in the RNA-induced Silencing Complex) and often leading to transcript degradation or translational inhibition (Zhang, 2009). RBPs
also direct localized expression/translation by recruiting transcripts to specific
subcellular sites, such as neuronal termini, where transcripts can be synthesized in
an activity-dependent manner without the need for long-range transport (Besse and
Ephrussi, 2008). RBPs are also involved in the editing of RNA (such as deamination
of adenosines to inosines; for a review, see Gott and Emeson, 2000): such changes in
the coding region can lead to the creation or abolition of start/stop codons or amino
acid substitutions (Maas et al., 1996), alter the protein sequence by changing splice- sites (Danecek et al., 2012; Rueter et al., 1999), or impact regulation by changing
UTR or intronic sequences. Finally, RBPs regulate the 3’ end processing and the adenylation state of mRNAs, thereby controlling their stability and degradation.
1.2. The CUG-BP, Elav-like family (CELF) of proteins
The CUG-BP, Elav-like family of proteins comprises a group of highly evolutionarily conserved RBPs (Barreau et al., 2006; Dasgupta and Ladd, 2012;
Good et al., 2000).
2 1.2.1. Nomenclature
CELF proteins have been studied for decades, in different species and for different biological functions. Understandably, this has resulted in a multitude of disparate names found in the literature. For clarity, all CELF proteins in this document will be referred to by their “CELF” annotation (CELF1 through CELF6), which is the most current system. A brief summary of the different CELF proteins and their many names is given in Table 1. An attempt was made to provide the protein symbols according to the naming conventions provided by the appropriate community standards (indicated in the notes for Table 1). In invertebrates, two
CELF proteins can be found, while in vertebrates these two proteins are each represented by subfamily consisting of multiple members (discussed in the next section). With some exceptions, six CELF members are found in vertebrates: CELF6 has not found in fish and frogs, while a deletion event has resulted in the loss of
CELF3 in the chicken genome. Notably, a genomic duplication in the ancestral lineage of the Xenopus has resulted in the polyploidy of all but one species in this genus: while the African Clawed Frog, Xenopus laevis, is pseudotetraploid (Amaya et al., 1998), X. tropicalis is diploid. Therefore, X. laevis has two sets of most genes (e.g.,
Celf1-a and Celf1-b are co-orthologs of the X. tropicalis Celf1). However, the duplication event was evolutionarily recent enough that duplicated genes are remarkably similar to each other and are considered functionally redundant. In
cases where a specific CELF ortholog is being referred to, the species-appropriate
notation will be used; for multiple orthologs, the human notation will be used.
3 Homo sapiens Mus musculus Gallus gallus Xenopus laevis Xenopus tropicalis Danio rerio elegans Caenorhabditis melanogaster Drosophila Species c
d
f
b h g e,# ,§
e
Brunol2 Eden - CUG CUG BRUNOL2, CUG Cugbp1, Eden - Cug CELF1 bp BRUNOL2 - - - - BP, CUG BP, CUG BP, CUG a, - bp1, Brul NAB50 Brunol2 - - - a BP1, BP1, BP1
b
p
Bru, Bru -
- ETR - b
1* Brunol3 2*
ETR - ETR - NAPOR, BRUNOL3 NAPOR, BRUNOL3 ETR -
Etr CELF2 3, CUG 3, CUG - - 3, CUG 3, Cug a
Etr3 Brunol3
- - - - BP2 BP2 BP2 bp2 % %
% , ,
- b
TNRC4, BRUNOL1, CAGH4 TNRC4, BRUNOL1, CAGH4 Brunol1 CELF3 - a Deleted Tnrc4
Etr - 1 Brunol1
- b
No previous UNC BRUNOL4 BRUNOL4 BRUNOL4 CELF4 Brunol4 Brunol4 name Bru - - 75, CELF**
3**
No previous BRUNOL5 BRUNOL5 BRUNOL5 CELF5 Brunol5 Brunol5 name
BRUNOL6 BRUNOL6 BRUNOL6 CELF6 Missing Missing Missing
4 Table 1. Names previously used for members of the CELF protein family in different species. The information in this table is based on the following sources: (Barreau et al., 2006; Dasgupta and Ladd, 2012; Good et al., 2000). aThe human protein symbol convention is used in column headings for clarity, but the species-specific conventions are used in the text Species-specific conventions were used based on the following sources: bFrom FlyBase (www.flybase.org St Pierre et al., 2014) cFrom WormBase (www.wormbase.org, release WS244) dFrom ZFIN (www.zfin.org Bradford et al., 2011) eFrom Xenbase (www.xenbase.org; Bowes et al., 2008; James-Zorn et al., 2013) f From Burt et al., 2009 gFrom MGI (www.informatics.jax.org; Eppig, 2006) hFrom HGNC (www.genenames.org; Gray et al., 2013) *A single gene in this species codes for a CELF1/CELF2 subfamily homolog **A single gene in this species codes for a CELF3-6 subfamily homolog #Due to a genome duplication event, X. laevis is pseudotetraploid (see text) %In early papers, CUG-BP2 referred to the hyperphosphorylated form of CELF1 (e.g., Roberts et al., 1997) §For a contrasting breakdown of the Drosophila orthologs, see (Delaunay et al., 2004)
5 1.2.2. Structure and RNA binding
As with members of their namesake Elav family of proteins, the CELF
proteins make use of RNA Recognition Motifs (RRMs) to bind to RNA substrates. The
standard RRM domain consists of a four-stranded anti-parallel β-sheet packed
against two α-helices, with RNA binding being coordinated along the β-sheet surface
and conserved residues in two conserved domains (RNP1 and RNP2) being used to
interact with the RNA (Tsuda et al., 2009). CELF proteins all have three conserved
RRMs, two (RRM1 and RRM2) in the amino terminal and one (RRM3) in the
carboxyl terminal, with a linker region (the ‘divergent domain’) located between
RRM2 and RRM3 (Ladd et al., 2001; see Figure 4). While the RRMs share high levels of similarity, both between paralogs (within species) and between orthologs
(between species), the ‘divergent domain’ is less well conserved. Within species, observed sequence similarities within this region have allowed for the definition of two subfamilies, consisting of CELF1-2 and CELF3-6 (Ladd et al., 2001; Ladd and
Cooper, 2004; see Figure 1).
The crystal structures of the CELF1 RRMs have been resolved and studied in
the presence of short oligonucleotides (Edwards et al., 2013; Teplova et al., 2010;
Tsuda et al., 2009), demonstrating that all three domains assume the same
β1α1β2β3α2β4 topology. Conserved residues in RRM1 and RRM2 are used to interact with the RNA, while RRM3 acts to augment the protein’s binding specificity.
CELF1 was initially hypothesized to bind to double-stranded RNA hairpins, a feature of the expanded CUG trinucleotide repreats that underlie pathology in the human
6
Figure 1. A dendrogram showing the evolutionary relationship between members of the CELF proteins family. Based on sequence alignments (Ladd et al., 2001; Ladd et al., 2004), the members of the CELF protein family can be subcategorized into the CELF1-2 subfamily and the CELF3-6 subfamily (see Table 1).
7 disease, Myotonic Dystrophy type 1 (DM1; OMIM#160900) (Timchenko et al.,
1996a). However, it was later shown that the protein binds to the single-stranded
sequences at the base of these hairpins (Edwards et al., 2013; Michalowski et al.,
1999). Sequence recognition has been extensively studied (summarized in Vlasova-
St Louis and Bohjanen, 2011). CELF proteins were shown to bind to GU-rich
elements (GREs; also referred to as embryonic deadenylation elements [EDENs]) by
RNA immunoprecipitation (RIP; Graindorge et al., 2008; Lee et al., 2010;
Rattenbacher et al., 2010; Vlasova and Bohjanen, 2008), systematic evolution of
ligands by exponential enrichment (SELEX; Faustino and Cooper, 2005; Marquis et
al., 2006), crosslinking immunoprecipitation (CLIP; Faustino and Cooper, 2005;
Marquis et al., 2006; Masuda et al., 2012), and surface plasmon resonance (SPR;
Mori et al., 2008), as well as X-ray crystallography and other approaches (Edwards et al., 2013; Teplova et al., 2010; Tsuda et al., 2009). The fly celf1 protein, Bruno, binds to sequences rich in AU dinucleotides (or Bruno responsive elements [BREs]),
which is also observed in CELF2 binding to AU-rich element (ARE) in the COX-2 3’
UTR during radiation-induced apoptosis in HeLa cells (Mukhopadhyay et al., 2003a)
and in human colon adenocarcinoma and mouse fibroblast cell lines (Sureban et al.,
2007). However, CELF1 binding to a similar ARE in the TNFα 3’ UTR was shown to
be mediated by binding to neighboring UGU-rich sequences (Zhang et al., 2008b),
and the majority of CELF protein binding is to GREs.
8 1.2.3. The ‘Divergent Domain’
The biological role of the ‘divergent domain’ has been less intensively
studied, but it has been shown to be important in subcellular localization (Fujimura
et al., 2008; Ladd and Cooper, 2004), trafficking (Moraes et al., 2013), RNA binding
(Takahashi et al., 2000), control of splicing regulatory activity (Han and Cooper,
2005; Singh et al., 2004), and enhancement of translation (Horb and Horb, 2010).
Finally, CELF1-mediated enhancement of deadenlyation is independent of RNA
binding, indicating another potential role for the ‘Divergent Domain’ (Paillard et al.,
2003).
1.2.4. Expression patterns and roles
The expression of CELF transcripts or proteins in embryonic tissues has been
shown in frogs (Amato et al., 2005; Wu et al., 2010b), zebrafish (Tahara et al., 2013),
chickens (Brimacombe and Ladd, 2007), mice (Karunakaran et al., 2013; Ladd et al.,
2001), and humans (Ladd et al., 2004; Lichtner et al., 2002; Meins et al., 2002).
These studies demonstrate that CELF1 and CELF2 are found in various tissues,
including striated muscle (both cardiac and skeletal), liver, and the nervous system,
among others. By contrast, CELF3-6 are largely restricted to the nervous system
(Brimacombe and Ladd, 2007; Ladd et al., 2004; Meins et al., 2002). In line with
these observations, deficiency of CELF4, both in human patients and in a mouse
model, results in seizures and defects of the eye (Halgren et al., 2012; Sun et al.,
2013; Wagnon et al., 2012; Wagnon et al., 2011; Yang et al., 2007). Similarly,
9 deficiency of Celf6 in a mouse model results in autism-related behaviors (Dougherty et al., 2013). However, other phenotypes suggest that low levels of protein in tissues outside the nervous system may still be important. For instance, mice deficient for
Celf3 exhibit defects of spermatogenesis (Dev et al., 2007), while knockdown and over-expression of Celf3 in the frog results in reduction and enhancement of proliferation in the gut and other endodermal-derived organs, respectively (Horb and Horb, 2010). There is currently no animal model for the study of CELF5.
Of the CELF1-2 subfamily, while CELF2 is expressed in a variety of tissues, it has been most intensively studied in the brain (Levers et al., 2002; Naha et al., 2009;
Otsuka et al., 2009; Pacini et al., 2005; Zhang et al., 2002; for review, see Ladd, 2012) and in cancer (Mukhopadhyay et al., 2003a; Natarajan et al., 2008). It has been shown to be involved in apoptosis following irradiation of cancer cells
(Mukhopadhyay et al., 2003b; Murmu et al., 2004; Pacini et al., 2005; but see Levers et al., 2002). Interestingly, however, its genomic deletion as part of a partial monosomy of the short arm of chromosome 10 has been suggested to be implicated in cardiac and thymus defects (Lichtner et al., 2002).
CELF1 has been the most studied member of the family and its activities and biological roles will be discussed in depth in the next few sections.
1.3. CELF1
CELF1 is found both in the cytoplasm and in the nucleus (Timchenko et al.,
1996a), suggesting multiple mechanisms of regulation.
10 1.3.1. Roles in the cytoplasm
During embryonic development in various animals, protein production is driven by maternal transcripts deposited into the egg during oocyte maturation. The precise activation and localization of these transcripts is important to various processes, including the patterning of the embryonic axes. In the fly, Celf1 (Bruno) binding to Oskar mRNA leads to the latter’s translational repression prior to its deposition in the posterior pole of the oocyte, and this repression is essential for the proper localization of this morphogen (Kim-Ha et al., 1995).
At the mid-blastula transition, maternal transcripts are rapidly degraded or translationally silenced, in favor of zygotic (embryonic) gene expression (Paris and
Philippe, 1990). In the frog, a GRE in the 3’ UTRs of maternal transcripts (EDEN) mediates the binding of Celf1 (EDEN-binding protein; Eden-bp) in frog oocytes, coordinating the deadenylation of target transcripts upon fertilization (Graindorge et al., 2008; Paillard et al., 1998; for a review, see Paillard and Osborne, 2003). This activity is facilitated by the presence of neighboring AREs in the frog c-mos 3’ UTR
(Audic et al., 1998). In mammals, the deadenylase was identified as the Poly(A)- specific ribonuclease deadenylase when TNFα and c-fos 3’ UTRs were found to be rapidly deadenylated using the same mechanism in HeLa cell extracts (Moraes et al.,
2006). CELF1 activity in promoting deadenylation was demonstrated to be evolutionarily conserved in showing that the frog EDEN can mediate deadenylation of reporter transcripts in fly oocytes, leading to translational repression (Ezzeddine et al., 2002); that the GRE in the mammalian c-jun 3’ UTR can mediate deadenylation
11 in the frog embryo (Paillard et al., 2002); and finally, that the human CELF1 can also mediate deadenylation (Paillard et al., 2003).
The ability of CELF1 to regulate the degradation of transcripts was investigated in different systems. A set of transcripts was shown to undergo rapid degradation in human T cells (Raghavan et al., 2002) and a computational analysis of the 3’ UTRs of these transcripts showed that they were enriched for GREs
(Rattenbacher et al., 2010). Furthermore, removal of CELF1 leads to the stabilization of these transcripts (Beisang et al., 2012; Vlasova and Bohjanen, 2008;
Vlasova-St Louis and Bohjanen, 2011). Similar investigations found transcripts in
HeLa cells (Rattenbacher et al., 2010; Vlasova et al., 2008) and in a C2C12 myoblast cell line (Lee et al., 2010) that are regulated by GRE-mediated degradation.
In addition to binding in the 3’ UTR, CELF1 was shown to enhance translation of p21 by binding to the 5’ UTR and antagonistically competing with calreticulin binding in fibroblasts (Iakova et al., 2004). Binding of CELF1 to CCAAT/enhancer
binding protein β (C/EBPβ) 5’ of the translation initiation site of the LIP isoform,
enhanced by Epithelial Growth Factor Receptor (EGFR) signaling, increases the
expression of the LIP isoform in mammary epithelial cells (Baldwin et al., 2004). The regulation of LIP translation was later demonstrated to represent CELF1-mediated enhancement of translation through interactions with eIF2α in liver following resection (Timchenko et al., 2005).
12 1.3.2. Roles in the nucleus
Myotonic Dystrophy (DM1) is a late onset muscular dystrophy genetically
characterized by the pathogenic expansion of CUG trinucleotide repeats in the 3’
UTR of the DM Protein Kinase (DMPK) gene. The human CELF1 was first identified
in search of a protein that would bind to CUG repeat RNA (and named CUG-Binding
Protein; CUG-BP) in HeLa cell extracts, with the majority of the binding found in the
cytoplasm (Timchenko et al., 1996b). However, the protein was also found to bind to
a nuclear polyadenylated RNA-binding protein, Nab2p, in a yeast-two-hybrid screen
in Saccharomyces cerevisiae (Anderson et al., 1993), and named hNAB50
(Timchenko et al., 1996a), suggesting a nuclear role. A nuclear role for CELF1 was further supported by the observation that CELF1 accumulated in the nucleus in cells from DM patients (Timchenko et al., 1996a). In the nucleus, CELF1 was found to regulate the alternative splicing of pre-mRNAs (Philips et al., 1998) and this function is conserved from humans to worms (Loria et al., 2003). The cardiac Troponin T
(TNNT2) gene (protein often written as cTnT) contains cis elements flanking exon 5 that mediate the tissue-specific inclusion of this exon in embryonic striated muscle
(Ryan and Cooper, 1996). Two of four of these Muscle-specific Splicing Enhancers
(MSEs) were found to contain GREs and CELF1 was shown to induce the inclusion of this alternative exon in DM1 patient cells, in which CELF1 accumulates in the nucleus (Philips et al., 1998). A similar study showed that CELF1 induce skipping of exon 11 of the Insulin Receptor (INSR) transcript (protein symbol: IR) through a binding site in intron 10 (Savkur et al., 2001). In addition to evaluating the endogenous TNNT2 or INSR transcripts, these studies made use of mini-genes, in
13 which an alternative exon flanked by test regulatory sequences (such as TNNT2 exon 5 flanked by MSEs from introns 4 and 5) are inserted between constitutively included exons and expressed in cells (Ryan and Cooper, 1996). Most studies of alternative splicing regulation of different targets by CELF proteins were carried out using similar reporters and care should be taken in extending these findings to endogenous targets in tissues and developmental stages of interest (for a compilation of many of these targets, see Dasgupta and Ladd, 2012 and Barreau et al., 2006). Using such mini-genes, the other CELF proteins were also shown to have alternative splicing regulatory capacity (Ladd et al., 2001; Ladd et al., 2004).
Nevertheless, the Philips and Savkur studies underscore the observation that
RBPs such as CELF proteins can regulate both inclusion and skipping events (Philips et al., 1998; Savkur et al., 2001). Later studies attempted to elucidate binding patterns that would mediate one event over another: a study of developmentally regulated alternative exons suggested that CELF1 binding motifs were predominantly found in the intron immediately downstream from the regulated exon, and their presence within the first 250 bases was associated with increased inclusion, while their presence within the last 250 bases was associated with reduced inclusion (Kalsotra et al., 2008). By contrast, a different analysis of alternative splicing events in a mouse myoblast cell line indicated that CELF1 binding to downstream introns associates with increased inclusion, while binding to upstream introns associates with increased skipping (Masuda et al., 2012). Both studies also highlighted the fact that binding motifs for different RBPs are commonly found in the same vicinity, suggesting that regulation in vivo is likely a
14 function of cooperative and antagonistic interactions between different factors
(Charlet-B. et al., 2002a; Gromak et al., 2003; Sureban et al., 2007).
1.3.3. Biological functions of CELF1
The mechanisms of regulation by CELF1 have been studied, but the biological functions of these regulatory events are less well understood. The significance of the regulation of maternal transcripts in frog embryos has been described, but the role of each target is not specifically known. The same can be said for the vast majority of validated and proposed targets of CELF1 regulation. One exception has been
mentioned above: the function of fly Celf1 repression of Oskar translation is
important to the latter’s localization and subsequent patterning (Kim-Ha et al.,
1995).
The mechanism by which CELF1 regulates p21 and C/EBPβ translation in
fibroblasts, mammary epithelial cells, and resected or old livers was described
above (Iakova et al., 2004; Timchenko et al., 2006; Timchenko et al., 2005),
indicating roles in normal biology and in pathology. In cells, this mechanism is
believed to allow CELF1 to impact both cell cycle arrest and proliferation, and this
involvement depends both on the phosphorylation state of CELF1 and on binding
competition with CRT, both mediated by post-transcriptional regulation (reviewed
in Barreau et al., 2006), which is discussed in the next section.
In other developmental systems, CELF1 was shown to have a role in the
development of both mesodermal and endodermal tissues. In C. elegans, the CELF1-
15 2 ortholog (Etr-1) is expressed in a muscle-specific manner and is essential for
muscle formation (Milne and Hodgkin, 1999), while the frog Celf1 was shown to be
involved in the formation of somites, which give rise to the skeletal musculature
(Cibois et al., 2013; Cibois et al., 2010; Gautier-Courteille et al., 2004). Finally,
studies of celf1 in zebrafish have indicated roles in symmetry patterning, as well as
cellular processes that underlie the development of endoderm-derived organs
(Matsui et al., 2012; Tahara et al., 2013). Interestingly, Celf3 was also shown to regulate endoderm proliferation and differentiation in frog embryos, through translational enhancement of cyclin A2 (Horb and Horb, 2010).
These regulatory mechanisms were also studied in disease. Aberrant splicing of some targets in patient myocytes (e.g., TNNT2 and INSR, as described above) has been used to explain some symptoms in DM1 pathology (Savkur et al., 2001), such as contractile defects and insulin resistance. The regulation of p21 and C/EBPβ by
CELF1 was shown to result in slowed differentiation of DM1 patient myoblasts
(Salisbury et al., 2008; Timchenko et al., 2001b), as well as in altered muscle fiber differentiation in mouse models of DM1 pathology (Timchenko et al., 2004). CELF1 regulation of alternative splicing is believed to play an important role in a developmental maturation switch in the splicing of many genes shortly after birth (a fetal-to-adult transition). In DM1 affected tissues, changes in CELF1 levels and
activity that recapitulate patterns of fetal and neonatal stages are believed to drive
reversal of this developmental switch and underlie DM1 pathology (Kalsotra et al.,
2008; Ladd et al., 2005a).
16 1.3.4. Mouse models for the study of CELF1
Since CELF1 is highly expressed in the placenta (Ladd et al., 2004), it is not surprising that mice in which CELF1 is systemically inactivated largely die perinatally (Cibois et al., 2012; Kress et al., 2007), likely due to placentation defects.
When the mutant allele is expressed on a mixed background, however, some mice expressing no detectable Celf1 survive and exhibit only growth retardation and spermatogenesis defects (Cibois et al., 2012; Kress et al., 2007). The effect of muscle- specific over-expression of Celf1 appears to be dose-dependent, as morbidity ranges from growth retardation with normal survival (Timchenko et al., 2004) to neonatal lethality (Ho et al., 2005). Heart-specific over-expression also appears to be dose- dependent, ranging from no phenotype to dilated cardiomyopathy and death within
14 days of transgene induction (Koshelev et al., 2010). Heart-specific repression of nuclear CELF activity by transgenic expression of a dominant negative CELF protein
(NLSCELFΔ; (Charlet-B. et al., 2002a)) suggests that the role of CELF proteins in the
heart may also be sex- (Ladd et al., 2005b) and developmental stage-specific
(Terenzi et al., 2009). These mice exhibit profound dilated cardiomyopathy, contractile defects, and activation of the Serum Response Factor (SRF) pathway
(Dasgupta et al., 2013).
1.4. Model organisms
Three model organisms are discussed in this work: mouse, chicken, and frog.
Each organism was chosen for its suitability for specific aspects of the studies
17 described. The model organisms are compared below based on the following criteria:
1. The facility of embryo collection and staging,
2. The facility of synchronized development and the number of embryos
available for study,
3. The timing of the developmental arc, and
4. The ability to manipulate the embryo.
1.4.1. The mouse
The benefits of the mouse as a model organism are well known, including the facility of genetic manipulation and the availability of a myriad of transgenic and mutant lines. However, as a developmental system, the mouse also presents challenges. The mouse embryo develops in utero and removal from the dam is necessary for precise staging. Litter sizes are relatively small (usually less than fourteen pups) and synchronization of multiple litters, by timed mating, is largely a matter of luck. At stages relevant to the studies described here, embryos are quite small (and hearts are miniscule) and the collection of a sufficient number of specimens, if tissues are to be used for certain common assays (such as western blot analysis), is both labor intensive and time consuming. Embryos were staged based on embryonic (E) days post coitus and Table 2 illustrates that mouse embryos undergo cardiac morphogenesis between days E8 and E14. In the studies reported
18
Mouse1 Chicken1 Frog1
(time) (time)
HH92 NF282 Straight heart tube E82 (~30 hrs)5 (~1 day)4
HH102 ~NF332 Contraction begins E8.52 (1.5 days)3 (~2 days)4
HH11-162 ~NF33-362 Looping E8.52 (1.8-2.5 days)3 (~2 days)4
HH16 Trabeculation NF414 (2.5 days)3
Endocardial HH172 ~NF412 E9.52 cushions form (2.6 days)3 (~3 days)4
HH24-28 Cushions remodel (4-5.6 days)3
HH26-30 Trabeculae coalesce (4.8-6.4 days)3
Atrial septum NF45 complete (~4 days)4
Atrioventricular HH35 No equivalent septum formed (8.7 days)3 structure
Morphogenesis HH35 NF46 E14 complete (8.7 days)3 (~4 days)4
Table 2. Comparative timeline of cardiac morphogenesis in mouse, chicken, and frog. 1Staging systems used: E, embryonic day post-coitus in mouse; HH, Hamburger & Hamilton in chicken; NF, Nieuwkoop & Faber in frog. 2From Ashe M, 2005 3From Wessels A, 2000 4From Faber., 1967 5From Hamburger and Hamilton, 1992
19 here, wild type embryos were collected at two representative stages for some
studies, while adult wild type and transgenic animals were used for others.
1.4.2. The chicken
The chicken has long been used as a model organism for heart development
and a large body of literature has accumulated on the avian heart. Like the
mammalian heart, the avian heart has four-chambers, and developmental processes such as patterning, induction, and morphogenesis are largely conserved between birds and mammals. Chicken embryos develop in ovo, allowing for the acquisition of large numbers of fertilized eggs and synchronized incubation for collection of dozens of embryos of the same stage per day. Also, embryonic chicken hearts are larger than mouse hearts of comparable stages. The developmental staging system of avian embryos (Hamburger and Hamilton [HH]; Hamburger and Hamilton, 1992) is well established and incubation periods necessary for different stages are well described. In the chicken embryo, cardiac morphogenesis is complete by embryonic day 8 (stage 35). A limitation of the chicken embryo is the difficulty of knocking down or over-expressing protein in developing embryos and the absence of transgenic animals. In the studies described here, the chicken embryo was used due to the large number of hearts that could be collected (on the same day) for pure primary cardiomyocyte isolation.
20 1.4.3. The frog
The frog has also been a traditional model organism in developmental studies,
although it has largely been used to investigate signaling and regulatory processes
in oocytes and in very early embryonic stages. Procedures for the laboratory
production and husbandry of frog embryos have been well established, and
hundreds of embryos can be easily produced from each female per day. Following
fertilization, the embryos are free living in a simple salt solution and can be quickly
and non-invasively staged using a well described staging system (Nieuwkoop and
Faber [NF]; Faber., 1967). A great strength of the frog embryo has been the ease
with which a variety of molecules, including nucleic acids and proteins, can be
microinjected into early-stage (2- to 4-cell stage) embryos. Such techniques have been extensively used for evaluating the effects of drug treatments, as well as gain- and loss-of-function of different proteins. Embryos develop rapidly and become progressively transparent, allowing for morphological and functional analyses, and cardiac morphogenesis is complete by day 4 post-fertilization (stage 46). For the studies described here, an important advantage of the frog embryo is the fact that its small size allows it to exchange oxygen with its environment in the absence of a beating heart, without negatively impacting the development of other tissues
(during stages relevant to these studies). Unlike in the chicken, powerful new
transgenic tools have recently been introduced for the Western Clawed Frog (X.
tropicalis), and these may prove to be quite useful in future studies.
21 1.4.4. Comparative staging
Equivalent stages of cardiac development are provided in Table 2 for the
three model organisms presented in this work (i.e., mouse, chicken, and frog). Note that comparisons between developmental stages are not absolute, as different organs develop at different rates. Table 2 only compares a small number of monumental events during cardiac morphogenesis and is designed to illustrate the differences in the developmental time courses of different species. Descriptions of these events are provided in Chapter 3.
1.5. The current work
The involvement of CELF1 has been extensively studied in disease (e.g.,
DM1), in different organ systems (e.g., brain, liver), and during different developmental transitions (e.g., frog oocytes and mouse post-natal maturation).
However, the role of CELF1 during normal embryonic heart development has been largely neglected. As discussed above, CELF1 is capable of regulating target transcripts in diverse ways, upregulating some while downregulating others, and the targets themselves may be gatekeepers to larger networks or biological pathways. The work presented in this document represents a set of investigations intended to elucidate the biological role of CELF1 during cardiac morphogenesis. In
Chapter 3, the current state of knowledge on the role of RBPs during heart development is described, including CELF proteins, and an overview of vertebrate cardiogenesis and cardiac morphogenesis is provided. In Chapter 4, the diversity of isoforms and expression patterns of CELF1 and CELF2 are explored in the
22 vertebrate embryo (i.e., chicken and mouse). These findings suggest tissue-specific
roles for these proteins, as different isoforms were found in different tissues and in
different subcellular compartments. In pursuit of direct targets of CELF1 regulation
in the embryonic heart, a previously unannotated exon is shown in Chapter 5 to be
a novel target of CELF1 regulation of alternative splicing. CELF1 levels are
correlated with skipping of this exon in MYH7B, and inclusion of this exon is
associated with lower transcript levels, suggesting that CELF1 can regulate the levels of MYH7B by regulating its splicing. This report also illustrates the regulatory
complexity that can be exercised by RBPs. Finally, CELF1 function was pursued, in
Chapter 6, by knocking down CELF1 both in cultured embryonic heart muscle cells
and in whole embryos. This study illustrates the involvement of CELF1 in the
organization of the contractile apparatus in cultured cardiomyocytes and in
embryonic hearts. CELF1 is shown to have a role in cardiomyocyte proliferation in
culture, and future studies will investigate this role in vivo. Finally, CELF1
expression is shown to be essential in the embryo for heart morphology and cardiac
looping, an important step in cardiac morphogenesis.
23 Chapter 2: Materials and Methods
2.1. Animal use
All animal work was carried out in accordance with the recommendations of
the American Veterinary Medical Association and in compliance with institutional
and federal guidelines for the ethical care and use of laboratory animals (protocol
numbers: 2011-0493, 2011-0495, 2011-0547, 2014-1201, ARC 07926, and ARC
08612). All efforts were made to minimize pain and distress during animal
husbandry and euthanasia.
2.2. Chickens
Fertilized White Leghorn Hy-line W-36 chicken eggs were purchased from
the Department of Animal Sciences at The Ohio State University or from Charles
River Laboratories. Chicken embryos up to embryonic day 14 were used in this study, which are not subject to federal regulation and do not require approval from the Cleveland Clinic Institutional Animal Care and Use Committee. Chicken embryos were euthanized by decapitation immediately upon removal from the egg, which is consistent with the recommendations of the American Veterinary Medical
Association Panel on Euthanasia for euthanasia of birds.
2.3. Chicken embryonic tissue and embryo collection
Fertilized chicken eggs were kept at 15°C (59°F) for up to one week until use.
Eggs were incubated at 100°F, 40-60% humidity with auto-turning until the
24 appropriate developmental stage was reached (stage 10 ≈ 1.5 days; stage 14 ≈ 2
days; stage 18 ≈ 3 days; stage 26 ≈ 5 days; stage 29 ≈ 6 days; stage 35 ≈ 8 days; and
day 14, where noted). At collection, embryos were extracted from the eggs and
staged using the Hamburger and Hamilton staging system (Hamburger and
Hamilton, 1992). Embryos were then either fixed for downstream applications or dissected for tissue collection.
2.4. Chicken embryonic primary cardiomyocyte isolation and culture
Fertilized eggs were incubated at 100°F, 40-60% humidity, with auto-turning for 8 days (stage 35). Embryos were then removed from the eggs and the hearts were dissected and collected as previously described (Ladd and Cooper, 2004), with minor modifications. Hearts were washed three times in Hank’s Balanced Salt
Solution (HBSS), minced using a straight edge blade in a culture dish, and resuspended in digestion solution (HBSS supplemented with 0.13% w/v trypsin
[TRL3; Worthington Biochemical], 0.13% w/v collagenase [CLS-2; Worthington
Biochemical], and 0.03% w/v DNase I [D; Worthington Biochemical]). Tissue was digested at 37 °C with rocking for three 20-minute incubations. After each incubation, Cardiomyocyte Growth Medium (Minimal Essential Medium [MEM] supplemented with 10% heat-inactivated horse serum, 1% Penecillin/Strepomycin,
5% chick embryo extract [day 12 chicks were collected, decapitated, and passed through a 60 mL syringe. An equal volume of Minimal Essential Medium was added to the homogenate and they were allowed to incubate at room temperature for two hours, with rocking. Suspensions were centrifuged at 500 xg for 5 min, the
25 supernatant was passed through a 0.45 µm filter, and the solution was stored at
−80 °C until use]) was added to the suspension, tissue was collected into a loose pellet by low-speed centrifugation (60 xg for 2 minutes), cells (in the supernatant) were removed, and the tissue was resuspended in digestion solution and allowed to continue digesting. The separated cells were pelleted (500 xg for 5 minutes) and resuspended in Cardiomyocyte Growth Medium on ice. Once all incubations were complete, the cells were collected by pelleting and resuspended in 1.082 g/mL
Percoll solution (in 1X Ads buffer: 116mM NaCl, 0.83mM MgSO4, 5.4mM KCl,
12.5mM NaH2PO4, 5.6mM dextrose, 20mM HEPES, 0.002% w/v Phenol Red, pH 7.3).
A Percoll gradient was formed by adding 4 mL 1.050 g/mL Percoll solution to a 15- mL conical tube, 4 mL 1.060 g/mL Percoll solution was added below the first layer, and the cell suspension was finally added to the bottom of the tube. The gradient was centrifuged at 2000 xg in a fixed angle rotor for 25 minutes to separate the cardiomyocytes. Cells were reclaimed from the interface between the 1.060 g/mL and the 1.082 g/mL layers, washed 3 times in 1X Ads buffer, resuspended in
Cardiomyocyte Growth Medium, counted, and plated (250,000 cells per plate in 35 mm plates; the same number of cells were plated on 60 mm plates for 5-ethynyl-2’- deoxyuridine [EdU] staining experiments); plates or coverslips (BD Biosciences) were coated with fibronectin (50 µg/mL working solution; Sigma) for at least 1 hour prior to plating. Cells were observed daily and medium was changed every 2-3 days.
2.5. Transfection of primary embryonic cardiomyocytes with siRNAs
Unless otherwise noted, cardiomyocytes were transfected 24 hours after
26 plating using Lipofectamine 2000 transfection reagent (Life Technologies) and a final concentration of 100 nM siRNA. Two siRNA duplexes (si1 and si2) against
CELF1 (Vajda et al., 2009), as well as siGLO Green and siGLO Red control siRNAs
(non-targeting transfection indicators; referred to in the text as “siCont”) were purchased from Dharmacon (now GE Healthcare). Unless otherwise noted, cells were collected by scraping 72 hpt. For western blot analysis, cells were collected into Protein Collection Buffer (0.2% Triton X-100 in PBS) supplemented with 1X cOmplete Protease Inhibitor Cocktail (Roche), sonicated, and stored at −80 °C until use. For real-time RT-PCR analysis, cells were collected into Trizol (Life
Technologies), sonicated, and stored at −80 °C until use. RNA was extracted as per the manufacturer’s instructions. Protein samples were quantitated using a Bio-Rad
Protein Assay. RNA samples were quantitated using a NanoDrop (Thermo). All experiments were repeated at least 3 independent times.
2.6. EdU staining of cultured primary cardiomyocytes
Cells were isolated as above, but plated at lower densities (250,000 cells per
60 mm plate). Plates were set up in duplicate (one for EdU staining and one for western blot analysis of knockdown efficiency). At 48 hpp, cells were pulsed with
EdU reagent (Click-iT EdU Alexa Fluor 488 Imaging Kit; Life Technologies) as per the manufacturer’s instructions. Briefly, the medium was replaced with fresh
Cardiomyocyte Growth Medium and EdU reagent was diluted in growth medium and added to the cells to a final concentration of 10 μM. Cells were returned to the incubator for 2 hrs before detection. To detect EdU incorporation, the cells were
27 removed from the incubator, washed briefly in PBS, and fixed in 4%
paraformaldehyde (PFA) for 15 minutes at room temperature (cells in duplicate
plates were collected for western blot analysis as described below). Cells were then
permeabilized in 0.2% Triton X-100 in PBS for 20 min at room temperature and
washed twice with 3% BSA in PBS. Click-iT reaction cocktail was added to the cells
and they were incubated for 30 min at room temperature (in the dark). Finally, cells
were washed with 3% BSA in PBS and VECTASHIELD hard-set mounting medium
with DAPI (Vector Labs) wash added to each plate. A coverslip was places into each
plate, the plate was allowed to dry for a few minutes, and then the coverslip was
sealed with nail polish. EdU (green channel) and DAPI (blue channel) were imaged
using an inverted epifluorescence microscope (Olympus) and positive cells were
counted.
2.7. Mice
Mice used for characterization of CELF1 and CELF2 expression patterns
(B6129F1; Taconic Farms, Inc.) were maintained in our colony. Transgenic mice
(lines MHC-CELF∆-10 and Myo-CELF∆-370) were maintained as hemizygotes, so wild type littermates were used for sex- and age-matched controls. Genotyping was
performed by PCR as previously described (Berger et al., 2011; Ladd et al., 2005b).
MHC-CELF∆-10 males were designated as “affected” or “unaffected” as previously
described (Dasgupta et al., 2013).
28 2.8. Mouse embryo collection
In order to obtain mouse embryos at specific stages, timed matings were
performed in our animal colony. At the appropriate day post-conception (day 11 or
14 for E11.5 or E14 embryos, respectively), pregnant females were euthanized by
CO2 inhalation followed by cervical dislocation. Embryos were removed, washed briefly in PBS, and fixed in 4% paraformaldehyde overnight at 4°C.
2.9. Western blotting and nuclear-cytoplasmic fractionation of tissues
For subsequent western blot analyses, tissues were collected on ice, snap- frozen in ethanol on dry-ice, and stored at −80°C until use. Upon use, tissues were homogenized in protein loading buffer (58 mM Tris pH 6.8, 9.3% glycerol, 1.86%
SDS, 1.86% β-mercaptoethanol, bromphenol blue) and sonicated. Protein concentrations were determined using a non-interfering (NI) protein assay (G-
Biosciences). Nuclear and cytoplasmic protein fractions were isolated from chicken tissues as previously described (Ladd et al., 2005a).
Protein samples were boiled briefly and subjected to SDS polyacrylamide gel electrophoresis (30 ug/lane). Proteins were transferred onto PVDF membranes and probed with primary antibodies (Table 3). Blots were exposed to Horse Radish
Peroxidase-conjugated secondary antibody and detected using Immobilon Western
Chemiluminescent HRP Substrate (Millipore). Three biological replicates were evaluated for each tissue, and each biological replicate consisted of tissue pooled from multiple embryos.
29
Primary antibodies: Antigen Host Application ID Distributor CELF1 Rabbit WB EPR8298(B) Epitomics CELF1 Mouse WB, IF 3B1 Santa Cruz Bio. CELF2 Rabbit WB 40-7500 Zymed Labs. ACTN2 Mouse IF EA53 Abcam TNNT2 Mouse IF O.N. 590 Abcam TNNT2 Mouse IF, IHC CT3d DSHBa TPM2 Rabbit IF ab11190 Abcam MYOM Mouse IF B4b DSHBa TTN Mouse IF T11 Sigma-Aldrich ACTA1 Rabbit IF NBP1-35265SS Novus Biologicals MEF2 Mouse WB C-21 Santa Cruz Bio. GAPDH Rabbit WB 6G5 Biogenesis, Inc. Meromyosin Mouse IF, IHC MF20c DSHBa β tubulin Mouse WB JDR.3B8 Sigma hnRNP-C1/C2 Mouse WB 4F4 Sigma Xpress tag Mouse WB R91025 Life Technologies
Secondary antibodies: Target Host Conjugate ID Distributor Mouse Sheep FITC 515-095-003 Jackson ImmunoResearch Rabbit Goat AlexaFluor488 A-11008 Life Technologies Rabbit Goat AlexaFluor594 A-11037 Life Technologies Mouse Goat HRP DC02L Calbiochem Rabbit Goat HRP 401393 Calbiochem
Table 3. Antibodies used in this work aAntibodies obtained from the Developmental Studies Hybridoma Bank were developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biology, Iowa City, IA 52242 bThis antibody was developed by Jean-Claude Perriard cThis antibody was developed by Donald A. Fischman, MD dThis antibody was developed by Jim Jung-Ching Lin
30 When appropriate, membranes were stripped using Restore Western Blot
Stripping Buffer (Thermo Scientific) and re-probed for GAPDH as previously
described (Dasgupta et al., 2013). Antibodies again nuclear (hnRNPC1/2) and
cytoplasmic (β tubulin) proteins were used to verify fraction purity. Equivalent
loading was confirmed both by GAPDH expression and Ponceau S staining for total
protein.
2.10. Immunofluorescence [IF]
Embryos were fixed in 4% PFA overnight at 4 °C. They were then transferred
to 20% sucrose and incubated at 4°C overnight. Embryos were then embedded in
2:1 20% sucrose:O.C.T. (Optimal Cutting Temperature solution; Electron Microscopy
Sciences), frozen on dry ice or liquid nitrogen, and stored at −80°C until use. Whole embryo sections (7-10 μm) were prepared using a cryostat microtome (Leica), post- fixed in ice-cold acetone for 10 min, and stored at −80°C until use. To probe for
CELF1, sections were permeabilized in 0.1% Triton X-100 in PBS for 20 min at room temperature, blocked in 3% BSA in PBS for 15 min at room temperature, and incubated with primary antibody (Table 3; with or without AlexaFluor586- conjugated phalloidin [Life Technologies]) in blocking solution at 4°C overnight (in a humidified chamber). Sections were washed in PBS, incubated with secondary antibody (Table 3; with or without DAPI) in blocking solution for at least one hour at room temperature (or overnight at 4°C) in a humidified dark chamber, washed in
PBS, and mounted using VECTASHIELD HardSet Mounting Medium (with or without
DAPI; Vector Laboratories).
31 Slides were sealed with nail polish stored at −20°C until use. Imaging
platforms used were: an inverted epifluorescence tiling microscope (DM5000B;
Leica Microsystems), using the ImagePro Plus software (Media Cybernetics), upright
epifluorescence microscopes (Olympus and Leica), and an upright laser confocal
microscope (Leica; using a water immersion objective lens). Images were processed
for publication using Adobe Photoshop CS2 (v.9.0.2), GIMP (v.2.8;
http://libguides.library.cofc.edu/gimp), and ImageJ (v.1.46; NIH; Schneider et al.,
2012; or Fiji v.2.0.0).
2.11. In situ hybridization
Chicken and mouse embryos were processed, sectioned, and hybridized as
previously described (LeMasters et al.). Antisense RNA probes against the chicken
CELF1 and CELF2 open reading frames were used, and sense probes for the same transcripts were used to confirm the absence of non-specific signal (Brimacombe and Ladd, 2007). Mouse Celf1 and Celf2 open reading frames were amplified from adult mouse brain RNA as previously described (Ladd et al., 2001). Amplicons were cloned into the pCR-Blunt II-TOPO vector using the Zero Blunt TOPO kit (Invitrogen) and confirmed by sequencing. Primers used for mouse Celf1 (5’-
ATGGCTGCGTTTAAGTTGG-3’ and 5’-TCAGTAGGGCTTACTATC-3’) and Celf2 (5’-
ATGTTTGAGCGCACTTCTG-3’ and 5’-TCAGTAAGGTTTGCTGTCG-3’) were ordered
from Integrated DNA Technologies (IDT). Sections were imaged on the LCN400
Slide Scanner platform (Leica Microsystems).
32 2.12. Semi-quantitative RT-PCR
Total RNA was extracted from Trizol (Life Technologies) according to
manufacturer’s protocols and all RNA samples were quantitated using a NanoDrop
(Thermo). Alternative splicing of primary embryonic cardiomyocytes was assessed
by semi-quantitative RT-PCR with 5’ radiolabeled primers as previously described
(Ladd et al., 2005b) using primers shown in Table 4 and conditions optimized for
amplification in the linear range with 100 ng total RNA per reaction: MYH7B = 64.6
°C annealing temperature, 20 cycles of amplification. MYH7B PCR products were resolved on 5% denaturing polyacrylamide:urea gels, scanned on a Storm 820
Molecular Imager, and quantified using ImageQuant software (GE Healthcare Life
Sciences). The identities of PCR products were confirmed by sequencing.
2.13. Real-time RT-PCR
Real-time analysis of transcript levels was carried out using either TaqMan
or Power SYBR Green reagents (Life Technologies), as indicated in Table 4. Total
RNA (1 μg) was reverse transcribed using the VILO kit (Life Technologies). The resulting cDNA was quantitated using the OligoGreen Quant-iT kit (Life
Technologies) as per the manufacturer’s instructions and stored at −20°C until use.
All primer sets and TaqMan probes were rigorously tested to confirm that their amplification curves were optimal (between 90% and 110% efficiency) and similar to each other before they were used (TaqMan probes were tested both singly and in duplex).
33
Gene Primersb or Probe ID Applicationa Species Amplicon (bp) CELF1 Gg03340922_m1 (FAM-labeled) qRT-PCR Chicken 67 celf1 F: ATGGAGGGCTGTTCCTCAC qRT-PCR Frog 95 R: ATTTGCTGCTGAAGTTGCTG CELF2 Gg03364304_m1 (FAM-labeled) qRT-PCR Chicken 65 celf2 F: GCTTGGAGGACTGACACCAC qRT-PCR Frog 71 R: ATGCACCCAGGTTACTGGAG GAPDH Gg03346982_m1 (VIC-labeled, qRT-PCR Chicken 107 primer limited) GAPDH F: GATACACAGAGGACCAGGTTG qRT-PCR Chicken 146 R: ACGGTTGCTGTATCCAAACTC Gapdh F: TCGTCCCGTAGACAAAATGG qRT-PCR Mouse 132 R: TTGAGGTCAATGAAGGGGTC GUSB Gg03358465_m1 (VIC-labeled, qRT-PCR Chicken 68 primer limited) MEF2A F: TGGAGGAGGTAATCTTGGAA Alt. splicing Chicken 126, 150 R: GGCTGCGTGGACTGAGAACT Mef2a F: ACCAGCCCTAATGCTTTGTCG qRT-PCR Mouse 196 R: GGTCTGTAGTGCTCAACATCCCAC MYH7B Gg03337745_m1 (FAM-labeled) qRT-PCR Chicken 67 MYH7B F: GGGAGGCTGCTGAATACCT Alt. splicing Chicken 291, 375 R: GCTTGAGGTTGTAGAGCACG Myh7b F: CCCGTTTCGACTTACTGGAG qRT-PCR Mouse 156 R: AGACCGGGAGCCATTTGTAT odc1 F: CCTCGATGGGTACAGATTTCG qRT-PCR Frog 147 R: GCTTCAGTCTCACTCCCAAG
Table 4. Primers and Probes aAlt. splicing= semi-quantitative RT-PCR for alternative splicing; qRT-PCR = real time RT- PCR for transcript levels bF = forward primer sequence, R = reverse primer sequence; all primers are shown 5’ à 3’
34 The StepOnePlus platform was used for all real-time RT-PCR experiments.
VIC-labeled TaqMan endogenous control probes and FAM-labeled TaqMan target
probes were obtained from Life Technologies and used in duplex. TaqMan reactions
consisted of 2 μL TaqMan Gene Expression Master Mix (Life Technologies), 1 μL
endogenous control probe, 0.25 μL target gene probe, and 5 ng cDNA. SYBR Green
reactions consisted of 10 uL Power SYBR Green Master Mix (Life Technologies), 150
pmol of each primer, and 5 ng cDNA.
All samples were run at least in triplicate (technical replicates) and at least
three biological replicates were included in each experiment. Data were analyzed by
the ΔΔCT method, normalizing target gene values to endogenous control (gapdh in the mouse; GAPDH or GUSB in the chicken; odc1 in the frog) values. Where developmental stages were compared, no endogenous control was found that did not also change during development, and so equal amount of cDNA template were used and ΔCT values were used for calculations instead. Where indicated, a method for scaling of data from multiple biological replicates was applied (Willems et al.,
2008). Unless otherwise indicated, error bars represent standard error of the mean
values for the biological replicates (the Willems method provides 95% confidence
intervals rather than standard error of the mean). Statistical comparisons of means were performed via one-tailed t-tests assuming unequal variances using Microsoft
Excel software. Differences were considered statistically significant when P ≤ 0.05.
35 2.14. Cross-linking immunoprecipitation (CLIP)
CLIP was performed using embryonic day 8 (stage 35) chicken hearts as
previously described (Ule et al., 2005) with a few modifications. In particular, due to
a high level of endogenous RNase activity in the heart, over-digestion of RNA was
observed without addition of exogenous RNase. Hence, appropriately sized CLIP
tags were obtained by addition of an RNase inhibitor. Immunoprecipitation was
performed using the 3B1 antibody (Santa Cruz), which specifically recognizes CELF1
with little to no cross-reactivity with other family members (Ladd et al., 2005a).
CLIP was performed following the general method of Ule and colleagues (Ule
et al., 2005), with appropriate optimization for our protein (CELF1) and tissue
(embryonic heart) at each step. Approximately 180-200 embryonic day 8 chicken
hearts were collected in 4 ml ice-cold PBS, gently triturated, and transferred on a
petri dish; the depth of the suspension was approximately 1 mm. The suspension
was irradiated three times for 400 mJ/cm2 in a Stratalinker (Stratagene model
1800) on ice, mixing between each irradiation. After irradiation, the suspension was immediately pelleted by centrifugation at 3000 rpm for 5 min at 4°C. Cross-linked pellets were frozen at −80°C until ready to use. Cross-linked lysate was resuspended in Buffer A (1X PBS, 0.1% SDS, 0.5% deoxycholate, 0.5% NP-40). Superase.In
(Ambion) was added to each tube to block RNA over-digestion. A “high RNase” sample treated with 1:100 RNase A (USB) was used as a marker. The samples were sonicated and each tube was treated with 30 µl RQ1DNase (Promega) for 5 min at
37 ˚C. These were ultracentrifuged for 20 min at 60,000 rpm at 4 ˚C, and supernatants were used for IP.
36 Dynabeads (Dynal) were equilibrated in Buffer A. Supernatant was incubated
with anti-CELF1 antibody (3B1, Santa Cruz) for 45 min at room temperature with
rotation, then washed three times with Buffer A and loaded onto the equilibrated
beads. Following a one hour incubation at 4˚C, the beads were washed with Buffer A followed by a high salt buffer B (5X PBS, 0.1% SDS, 0.5% deoxycholate and 0.5% NP-
40), and finally twice with 1X PNK buffer (50 mM Tris-Cl, pH7.4, 10mM MgCl2, 0.5%
NP-40). An on-bead CIP (Roche) treatment was performed. The beads were incubated for 10 min at 37 ˚C at 1000 rpm, then washed with 1X PNK-EGTA buffer
(50 mM Tris-Cl, pH7.4, 20mM EGTA, 0.5% NP-40) followed by 1X PNK buffer.
The 3’ linker was ligated on bead with T4 RNA ligase overnight at 16˚C. The following day the beads were washed 9-10 times with 1X PNK buffer, then treated with T4 PNK enzyme (New England Biolabs). Beads were washed three times with
1X PNK buffer. 40 µl of 1X PNK buffer and 40 µl Nu-PAGE LDS sample buffer (Life technologies) were added to the beads and boiled for 5 min. 1 µl β-mercaptoethanol was added to every 40 µl of the sample and boiled again. 40 µl supernatant from each tube was loaded onto a Novex NuPAGE 10% Bis-tris SDS-PAGE gel and transferred onto a nitrocellulose membrane. Smeared bands in the range of 70-120 kDa were cut out into a single eppendorf. This was digested with proteinase-K in PK buffer (100 mM Tris-Cl, pH 7.5, 50 mM NaCl, 10 mM EDTA) for 20 min at 37˚C at
1000 rpm. This incubation was repeated after addition of 200 µl of PK buffer in 7 M urea. RNA was extracted with phenol:chloroform followed by a 1:1 ethanol:isopropanol precipitation.
37 The 5’ linker was ligated to the resuspended RNA with T4 RNA ligase for 4
hours at 16˚C. DNA was digested with RQ1 DNAse, and RNA was re-isolated by phenol:chloroform extraction and ethanol:isopropanol precipitation. Reverse transcription was performed with P3 primer using Superscript III (Invitrogen). PCR was performed using P3/P5 primers and Accuprime Pfx Supermix with the following program: 95 ˚C for 5 min, followed by 35 cycles of 95˚C for 20 sec/61˚C for
30 sec/68˚C for 20 sec, and a final extension of 68˚C for 5 min. Samples were run on a 10% denaturing polyacrylamide gel, and DNA fragments in the range from 70-110 bp were extracted using a QIAEX II kit (Qiagen). The isolated PCR products were
amplified two more times using P3/P5 primers and Accuprime Pfx supermix. The products were desalted with GE G25 microspin columns (Amersham) and subcloned into a TOPO vector (Invitrogen). Although 384 clones were sequenced, a total of 736
sequences representing 564 unique tags were obtained due to concatamerization of
the tags. CLIP tags were identified by alignment to the chick (galGal3) genome by
BLAT search using the University of California, Santa Cruz (UCSC) genome browser
(Kent, 2002) and/or BLAST search of the NCBI database (Altschul et al., 1997).
2.15. Analysis of CLIP tags
CLIP tags were extracted from clone sequences using an in-house script
(Supplemental Script 1; in Appendix) that identified sequence tags bound by CLIP
adapters in the correct orientation, removed adapter sequences, and tabulated the
resulting tags. The nucleotide composition of the tags was analyzed using an in-
house script (Supplemental Script 2; in Appendix) that counted the occurrence of
38 each nucleotide combination (1 to 6-mer) in the tag dataset; this program made use
of a script posted by Mike Golvach, 2008, shared under the Creative Commons
Attribution 3.0, which generated oligonucleotide permutations. To characterize CLIP tag-containing introns, intron coordinates within annotated genes were downloaded from the UCSC genome browser (Kent, 2002) using the Galaxy platform
(Giardine et al., 2005), and compared to the genomic coordinates of the tags using an in-house script (Supplemental Script 3; in Appendix). The same approach was used to analyze CLIP data for CELF1 (Daughters et al., 2009) and Nova (Ule et al.,
2003) in postnatal mouse brain using the mouse mm9/NCBI37 genome and annotation (July 2007).
2.16. Frog embryo production
Female frogs (Xenopus laevis) were induced to produce eggs by intramuscular injection of human chorionic gonadotropin the day before eggs were to be collected. On the day fertilizations were performed, adult frogs were transported from the animal facility and transferred to 1X High-Salt Barth’s Solution
(110 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 1.7 mM MgSO47H2O, 488 µM
Ca(NO3)24H2O, 408 µM CaCl22H2O, 10 mM HEPES, pH 7.6). A male frog was
euthanized using MS222 followed by pithing, and testes were removed. and minced
in Steinberg Solution (58 mM NaCl, 671 µM KCl, 339 µM Ca(NO3)24H2O, 811 µM
MgSO47H2O, 3.81 mM Tris-HcL, pH 7.35-7.45). Eggs were collected into glass Petri
dishes on the bench, mixed with minced frog testis, and incubated in 0.1X Barth’s
Solution (8.9 mM NaCl, 102 µM KCl, 238 µM NaHCO3, 166 µM MgSO4, 48.8 µM
39 Ca(NO3)24H2O, 40.8 µM CaCl22H2O, 998 µM HEPES, pH 7.6, supplemented with
Pen/Strep) at room temperature for 30 minutes. Fertilization was initiated by immersion of the eggs and sperm in cysteine solution (114 mM, pH 7.8) for 15 minutes, with rocking on an orbital shaker. Fertilized eggs were washed three times with water and three times with 0.1X Barth’s Solution, and then allowed to develop normally on the bench. Embryos were staged using the Nieuwkoop and Faber staging system (Faber., 1967) and collected into different fixatives, depending on the application (see below).
2.17. Frog embryo morpholino oligonucleotide (MO) microinjection
All MOs were purchased from GeneTools. Celf1-targeting MO sequences were ordered as previously described (Gautier-Courteille et al., 2004): a mixture of two
MOs was used, one targeting Celf1a and the other targeting Celf1b. A control MO
(STD-MO) was ordered as offered by GeneTools. MOs were resuspended in distilled water to 1 mM and stored at 4 °C. Working solutions were prepared by diluting stocks in distilled water to 200 µM and solutions were stored at 4 °C until use.
Before use, working MOs were heated at 55 °C for 5 min, spun down, and kept on ice.
Eggs were collected and fertilized as described above. At the 2- to 4-cell stage, embryos were microinjected with MOs. MOs were injected into all blastomeres, using a pulled glass microinjection needle (4 nL/blastomere at the 2-cell stage or 8 nL/blastomere at the 4-cell stage; total 6.4 pmol/embryo [Gautier-Courteille et al.,
2004]). Embryos were transferred to 60 mm dishes containing solidified 2% agarose (prepared in 0.1X Barth’s Solution) and injected in 1X Barth’s Solution.
40 Embryos were kept in this solution for 1-2 hrs following injections before being transferred to fresh 0.1X Barth’s Solution. Fresh Barth’s Solution was replaced every day until the embryos were collected. Each experiment included Celf1-MOs-injected embryos, STD-MO-injected embryos, and uninjected embryos.
2.18. Whole-mount immunohistochemistry [IHC] and IF
Frog embryos were collected into Dent’s Solution (20% dimethyl sulfoxide in methanol; for IHC) or 4% PFA (for IF) at the appropriate stage and incubated at 4 °C overnight, with gentle rocking (Note: Dent’s fixation resulted in complete loss of
Phalloidin staining). Dent’s-fixed embryos were washed twice in methanol before being transferred into PBSw (0.1% Tween 20 in PBS) through a graded series of methanol-PBSw (75%, 50%, 25%; 5 minutes each). PFA-fixed embryos were washed briefly in PBS and then transferred directly into PBSw. Embryos were incubated in PBSw for 2 X 1 hr at room temperature, and then incubated in hybridization solution (3% BSA in PBSw) for 2 hrs at 4 °C. Primary antibody was added to the embryos (1:100; Table 3) and they were incubated overnight at 4 °C, with rocking. Embryos were washed 4 X 1 hr in PBSw, 1 X 2 hrs in hybridization solution, and then secondary antibody was added (1:1000; Table 3) and the embryos were incubated at 4 °C overnight, with rocking (for IF, embryos were incubated in the dark). Finally, embryos were washed 5 X 1 hr with PBSw. Embryos stained for IHC were treated using the DAB-Plus Substrate Kit (Life Technologies) for 5-30 min and then stored in PBS at 4 °C. Embryos treated for IF were cryo- embedded (as described above) and cryosectioned (several embryos were imaged
41 by whole-mount confocal microscopy: these were adhered to a culture dish using cyanoacrylate adhesive, overlaid with water, and imaged using a water-immersion objective).
2.19. Bright-field imaging
Bright-field images of injected embryos were acquired with a MicroPublisher
5.4 RTV camera (Q-Imaging) mounted onto a Leica dissecting microscope, using the
QCapture Pro (v.5.1.1.14) software.
2.20. Frog construct design
To create the pCS2-celf1a construct, the frog celf1a sequence was amplified from cDNA generated from stage 45 isolated hearts. The full-length transcript was amplified using a high-fidelity polymerase (Pfx50; Life Technologies), inserted into the pCR-Blunt II-TOPO backbone using the Zero Blunt TOPO PCR Cloning Kit (Life
Technologies), and transformed into TOP10 cells. The resulting clones were confirmed by sequencing of the full insert and compared to the reference (Xenopus tropicalis) sequence (NM_001090727.1): compared to the X. tropicalis sequence, the cloned sequence contained an inserted stretch within the linker between RRM1 and
RRM2 (that is predicted to result in a change at position p.104 from an A to an evolutionarily conserved G, as well as an insertion of 13 amino acids within the linker), an insertion within the variable domain that has been reported previously in various species (the “LYLQ” variant; Takahashi et al., 2000; Takahashi et al., 2001),
42 and one synonymous substitution (c.G1299C; p.G433G) within RRM3. The celf1a
insert was removed from the plasmid using EcoRI (New England Biolabs), ligated
into an EcoRI-linearized pCS2+ plasmid with T4 DNA Ligase (New England Biolabs), and transformed into α-Select Gold cells (Bioline). Insert orientation was confirmed by diagnostic digests and the final pCS2-celf1a plasmid was sequence-confirmed.
The pCS2-RTB33.51 construct was created by amplifying the RTB33.51 mini-gene
(Ryan and Cooper, 1996) out of the KS+ backbone using Pfx (Life Technologies) and primers designed for directional cloning (BamHI-forward primer and XhoI-reverse primer), and blunt-end ligating the amplicon into pCR4Blunt-TOPO. Clones were confirmed by diagnostic digests using PstI (New England Biolabs) and by sequencing. The insert was isolated by digestion with BamHI/XhoI (New England
Biolabs), ligated into linearized pCS2+ backbone with T4 DNA Ligase (New England
Biolabs), and transformed into α-Select Gold cells (Bioline).
2.21. Alternative Splicing Assay
Cultured COSM6 (kindly provided by Dr. Thomas Cooper, Baylor College of
Medicine) cells were grown in COSM6 Growth Medium (DMEM supplemented with
10% FBS, 2 mM L-glutamine, 1% Pen/Strep) in 60 mm culture dishes. When cells reached 50-60% confluence, they were transfected with plasmid/s using
Lipofectamine 2000 (Life Technologies). All cells were transfected with the pCS2-
RTB33.51 mini-gene (100 ng/plate) and half the plates were co-transfected with frog celf1a (pCS2-celf1a; 1 µg/plate). Cells were collected into Trizol (for RNA isolation) or Protein Loading Buffer (for western blot analysis) when they reached
43 90-100% confluence. All treatments were carried out in duplicate plates (one for protein and one for RNA) and all experiments were carried out at least 3 separate times. For western blot analysis, equal volumes were loaded per lane (cell densities were visually confirmed to be equivalent in all plates before collection). The mini- gene was reverse transcribed and amplified as previously described (Ladd et al.,
2001), using primers designed against the shared flanking constitutive exons
(forward primer: 5’-AGGTGCTGCCGCCGGGCGGTGGCTG-3’; reverse primer: 5’-
CATTCACCACATTGGTGTGC-3’) with a small amount of 5’-radiolabeled forward primer. Amplification products were separated on 5% polyacrylamide gels, scanned on a Storm 820 Molecular Imager, and quantified using ImageQuant software (GE
Healthcare Life Sciences).
44 Chapter 3: RNA binding proteins in the regulation of heart
development
This chapter has been adapted from published work: Blech-Hermoni Y and
Ladd AN (2013) RNA binding proteins in the regulation of heart development. Int J
Biochem Cell Biol 45: 2467-2478.
3.1. Introduction
Growing interest in the functional repertoire of RNA binding proteins (RBPs) has emerged as their potential to regulate gene expression has become more broadly appreciated. While the old paradigm of gene expression focused on the activation of transcriptional programs by DNA binding proteins, the roles of RBPs in post-transcriptional regulation have recently been given greater scrutiny. Post- transcriptional regulatory mechanisms have been identified at all levels of the life cycle of a transcript: regulation of pre-mRNA alternative splicing (Kelemen et al.,
2013), mRNA editing (Chateigner-Boutin and Small, 2011), transcript stability
(Schoenberg and Maquat, 2012), transcript localization (transport and
sequestration) (Medioni et al., 2012), and regulation of translation (Kong and Lasko,
2012). RBPs are involved in the regulation of each of these processes [for an
overview see (Glisovic et al., 2008)]; some specific examples are illustrated in
Figure 2. Some of these functions are nuclear, while others are cytoplasmic, or take place within other organelles such as mitochondria. There are several different types of RNA binding domains, which divide RBPs into structurally distinct families.
While many employ beta sheets as interaction surfaces to interface with client
45
Figure 2. Mechanisms of RBP-mediated post-transcriptional regulation. Schematic representations of mechanisms by which a number of proteins described in this review have been shown to regulate gene expression. Note that these are provided as examples; an exhaustive survey of RBP-mediated regulatory mechanisms is beyond the scope of this review. (1) RBFOX proteins regulate a variety of alternative splicing events by binding within introns flanking alternative exons. Binding upstream of an exon leads to skipping of that exon, while binding downstream of an exon leads to its inclusion (De Craene and Berx, 2013). (2) CELF2 directs the editing of a cytidine in the Apob transcript by binding to an AU-rich sequence element upstream of the editing site and recruiting ACF, a component of the editing machinery (Anant et al., 2001). (3) MBNL2 regulates the transport and localization of the Itga2 transcript to the plasma membrane by binding to a zipcode sequence in the 3’ UTR of the transcript (Adereth et al., 2005). (4) Multiple mechanisms have been proposed for how AUF1 regulates the stability of target transcripts, including the recruitment of the PARN deadenylase, leading to loss of the poly-A tail and rapid degradation of the RNA (White et al., 2013). (5) CELF1 enhances translation of the p21 transcript by antagonizing a regulatory protein, CRT, which normally blocks ribosome loading (Iakova et al., 2004).
46 transcripts and utilize aromatic residues and base stacking interactions to achieve recognition of targets, a detailed understanding of the binding properties of many
RBPs remain to be elucidated (Lunde et al., 2007). While their RNA binding domains can help catalog these proteins, RBP families are not characterized by unified functions. Individual RBPs may perform several functions within the same cell, and may have different functions in different cell types. The ability of RBPs to circumvent the transcription machinery allows them to quickly and selectively fine- tune expression, and this capacity has been recognized as especially important in developmental and pathological systems (Masuda et al., 2009; Misquitta et al., 2001;
Siomi and Dreyfuss, 1997). For example, during early zygotic development, when maternal transcripts are translated but the transcription machinery is silent, RBPs provide robust mechanisms for regulating gene expression to direct processes such as pattern formation and cell-type specification (Lee and Schedl, 2006).
The post-transcriptional regulation of gene expression by RBPs during development also has evolutionary consequences. Because of their generally small size and ability to rely on diffusion for tissue oxygenation, many invertebrate species lack a defined circulatory system. The invertebrate heart is typically a simple, beating tube or sac that moves fluid through the body via peristaltic contractions. Vertebrates, on the other hand, have closed circulatory systems with multi-chambered hearts. It has been proposed that vertebrates exhibit a greater degree of cellular and organismal complexity than invertebrates due in large part to expansion of the transcriptome (without proportional expansion of the genome) via an increase in alternative RNA processing, particularly pre-mRNA alternative
47 splicing (Ast, 2004; Maniatis and Tasic, 2002). Consistent with this, several RBP
families involved in alternative splicing regulation are differentially expanded in
vertebrates compared to invertebrates, whereas basal splicing machinery proteins
are generally invariant among all eukaryotes (Barbosa-Morais et al., 2006; Pascual et al., 2006).
The development of the heart is a complex and finely orchestrated process.
RBPs have been shown to be involved in nearly every step of heart development, from the establishment of cardiac lineages to the maturation of the heart after birth
(Figure 3). In addition, there are RBPs that are known to be expressed in the heart at some stage, but whose molecular and biological roles in heart development have not yet been determined. This review will focus on the expression and functions of
RBPs that have been implicated in the formation, morphogenesis, and maturation of the heart (Figure 4). It should also be noted that a set of RBPs known as Argonaute proteins interact with microRNAs (miRNAs) within the RNA-induced silencing complex (RISC) to destabilize or inhibit the translation of target mRNAs that possess sequence complementarity (Amiel et al., 2012). Although miRNA-mediated gene expression changes are important in the heart during normal development and disease (Chen and Wang, 2012; Ono et al., 2011), we will not discuss Argonaute proteins further in this review, as it is generally the miRNAs rather than their associated RBPs that are developmentally regulated.
3.2. Cardiac cell fate, heart tube formation, and differentiation
The heart is the first functional organ to form in the vertebrate embryo.
48
Figure 3. RNA binding proteins have been implicated in the formation, morphogenesis, and maturation of the heart. The primitive heart tube forms from precardiac mesoderm within the cardiac crescent. The heart tube undergoes extensive morphogenesis, including cardiac looping, endocardial cushion formation and remodeling, and myocardial trabeculation and compaction. Although the architecture of the heart is established during embryogenesis, maturation of the heart continues through postnatal life. RNA binding proteins that have been implicated in specific steps of heart development are indicated in blue. Abbreviations: OFT, outflow tract; RV, right ventricle; LV, left ventricle; SA, sinoatrial segment; atr, common atrium; AVC, atrioventricular canal; LA, left atrium; RA, right atrium; vc, vena cava; ao, aorta; pa, pulmonary artery.
49
50 Figure 4. Domain structure of RNA binding proteins implicated in regulation of heart development. Schematic representations of the type and position of important domains within the RNA binding proteins described in this review are shown. RNA binding domains and other domains characteristic of these RNA binding protein families are shown in color; other conserved domains are shown in black and white. Proteins and domains are not drawn to scale. Molecular functions of these RNA binding proteins within the heart are indicated, if known; additional functions of these proteins that have been demonstrated in other tissue types are not shown. The domain structures of these proteins are highly conserved across vertebrate species, and the depicted structures represent all homologs described in the text. Note that the structure of QK/How is conserved from human to fly, whereas to date CHAMP/Csm has only been described in mammals. All protein symbols are elaborated in the List of Abbreviations section.
51 Induction of cardiogenesis begins in the pregastrula embryo (Ladd et al., 1998;
Yatskievych et al., 1997), and precardiac cells become specified within the nascent
mesoderm as it arises during gastrulation (Brand, 2003). As presumptive
cardiogenic cells migrate out of the primitive streak, the heart-forming regions are
found on either side of the midline. Both positive and negative signals from nearby
tissues act to induce cells in the heart-forming regions to migrate rostrally and
medially to form the cardiac crescent (i.e., the primary heart field). A second cohort
of cells (i.e., the secondary heart field) arising from splanchnic mesoderm at the
arterial pole also contributes to the formation of the heart (Brand, 2003;
Dunwoodie, 2007). Cell migration, along with flexion and folding of the embryo, bring the heart-forming regions together at the midline where they fuse to form the primitive heart tube: an outer, myocardial layer surrounding an inner, endocardial layer. The linear heart tube is arranged from the sinoatrial segment (inflow tract) at the posterior end to the presumptive left and right ventricles, and finally the conus arteriosus (outflow tract) at the anterior end. Completion of the heart tube coincides with the assembly of nascent myofibrils in the cardiac myocytes, and beating begins.
In vertebrates, the primitive heart tube is later transformed into a multi-chambered
heart.
The inductive signals that establish cardiac cell fates have been long studied,
and include Wnt-β catenin, fibroblast growth factor (FGF) and Hedgehog proteins,
and members of the transforming growth factor β (TGFβ) superfamily such as the
bone morphogenetic proteins (BMPs) (Brand, 2003; Dunwoodie, 2007). These growth factors initiate signaling cascades that activate important cardiogenic
52 transcription factors of the NKX, TBX, GATA, and MEF2 families (Brand, 2003;
Dunwoodie, 2007). Much less is known about the contributions of post- transcriptional programs to the establishment of cardiogenic cell lineages, but several RBPs have been shown to participate in heart tube formation or cardiac cell differentiation (Figure 3).
3.2.1. Formation of the heart tube
Unlike the multi-chambered vertebrate heart, the heart of the fruit fly
Drosophila melanogaster remains a simple linear tube. The Drosophila heart tube is subdivided into an anterior “aorta” and a larger posterior cavity. Although the fly heart is structurally much simpler, gene expression programs important for cardiogenesis are largely conserved from flies to man. This conservation, along with
the ease of performing large-scale genetic screens, has made Drosophila a powerful
model system for the identification of important regulatory genes. For example, the
importance of NK-class homeodomain-containing transcription factors (such as
Nkx-2.5) in heart formation was first uncovered in studies of the fly paralog, tinman
(Bodmer, 1993). Similarly, RBPs have been identified in flies that are important in
the developing heart.
Held out wings (How) is a heterogeneous nuclear ribonucleoprotein (hnRNP)
K-homology (KH)-domain protein (Figure 4) involved in muscle function
(Baehrecke, 1997). It is most similar to the mouse Quaking (QK) and Caenorhabditis
elegans GLD1 proteins, which belong to the signal transduction and activation of
53 RNA (STAR) subfamily of KH-domain proteins. Members of this subfamily have been described to link signal transduction pathways to RNA metabolism in different tissues (Lasko, 2003). In flies, MAPK/ERK-dependent phosphorylation of How promotes its RNA binding activity, and How is phosphorylated in vivo in embryonic cardioblasts (Nir et al., 2012). A maternal how transcript is found broadly in the
Drosophila zygote, although no detectable protein product is detected (Zaffran et al.,
1997). A zygotic transcript is activated in later stages in the presumptive mesoderm, and then in cells of myogenic lineages and in epidermal muscle attachment cells
(Baehrecke, 1997; Zaffran et al., 1997). Later in development, how transcripts are restricted to the heart and to muscle attachment sites. This expression pattern is maintained during and following morphogenesis, with the addition of expression in adult muscle precursor cells attached to the wing imaginal discs, resulting in the phenotype for which the gene is named (Baehrecke, 1997; Zaffran et al., 1997).
Despite this broad early expression, histological analysis of mutants suggests that perturbation of how expression does not result in disruption of striated muscle formation, but rather leads to functional aberrations including reduced cardiac beat rate and weakened contraction (Zaffran et al., 1997). A recent study has suggested that How regulates the expression of sarcomeric proteins, although it is unclear whether this relationship is direct or indirect (Nir et al., 2012). How has also been shown to be part of a genetic pathway with the extracellular matrix protein Slit, its receptor Robo, and dystroglycan that controls formation of the cardiac lumen during heart tube morphogenesis (Medioni et al., 2008).
54 The mouse QK has nuclear and cytoplasmic isoforms generated by
alternative splicing (Kondo et al., 1999). QK binds to consensus UACU(C/A)A hexanucleotide sequences (Ryder and Williamson, 2004), and regulates the alternative splicing of transcripts encoding proteins involved in myelination in the adult brain (Wu et al., 2002). An allelic series of mice bearing different Qk mutations, many of which are embryonic lethal, has revealed roles for QK in formation of the heart and vasculature (Justice and Hirschi, 2010). The specific targets of QK, and the
specific functions of different QK isoforms, within the developing heart have yet to
be elucidated. Another related STAR protein, Sam68, is also found in mouse heart,
among a variety of other tissues, but its contributions to cardiac development or function have not been studied (Richard et al., 2005).
Heart and RRM expressed sequence (hermes), the homolog of the human
RNA binding protein with multiple splicing type 1 (RBP-MS type 1), is an RNA
recognition motif (RRM)-containing RBP (Figure 4) expressed in a variety of
embryonic tissues including heart (Gerber et al., 1999). In Xenopus laevis, hermes is
first detected in the developing heart concomitant with the first cardiac
differentiation markers shortly before the coalescence of the linear heart tube, and
remains high throughout morphogenesis (Gerber et al., 2002; Gerber et al., 1999).
Likewise, in the chicken embryo HERMES is first detected in a crescent of Nkx-2.5-
positive cardiac precursor cells, before expanding to encompass the entire
myocardial layer of the heart tube (Wilmore et al., 2005). At later stages, however,
its expression is curtailed in the outflow tract and persists primarily in the
ventricles and atria (Wilmore et al., 2005). In the mouse, Hermes expression is
55 absent in the early heart tube, but becomes widespread following looping, after
which its expression becomes increasingly atrial (Gerber et al., 1999). Hermes likely regulates cardiac cell fate, as over-expression in the Xenopus embryo leads to
dramatic reductions in the expression of cardiac markers, including cardiac α-actin
and Nkx-2.5, and failure of heart tube formation (Gerber et al., 2002). This effect
seems to be specific to the role of hermes in heart, since over-expression in somites
does not result in aberrant skeletal muscle gene expression.
3.2.2. RBPs in cardiomyocyte differentiation and myofibril development
Recent studies investigating transcriptome-wide changes in alternative
splicing during differentiation of pluripotent stem cells have highlighted the
importance of post-transcriptional RNA processing in cell fate decisions and
differentiation (Brandenberger et al., 2004; Cloonan et al., 2008; Pritsker et al.,
2005; Salomonis et al., 2009; Salomonis et al., 2010; Yeo et al., 2007). In a
comparison of human embryonic stem cells before and after differentiation into
either cardiac or neural progenitors, Salomonis and colleagues identified alternative
splicing events common to both differentiation pathways, but also events that were
specific for the cardiac lineage (Salomonis et al., 2009). To what extent these alternative splicing events direct cardiomyocyte differentiation, and the RBPs responsible for their regulation, however, remain unknown. Knockdown/knockout experiments in fish, frogs, and rodents have identified several RBPs that are
56 important during development for cardiomyocyte-specific gene expression,
myofibril assembly, and myocardial function in vivo (Figure 3).
RNA binding motif (RBM) proteins are a subgroup of loosely related RRM- containing proteins (Figure 4) with varying functions, including splice site selection and nonsense-mediated RNA decay (Sutherland et al., 2005). RBM24 is up-regulated in both human and mouse embryonic stem cells upon differentiation into cardiomyocytes (Miller et al., 2008; Xu et al., 2009). Orthologs of RBM24 are expressed first in cardiac precursors, and then in the differentiated myocardium in zebrafish, frog, and mouse embryos (Fetka et al., 2000; Maragh et al., 2011; Miller et al., 2008; Xu et al., 2009). Knockdown of rbm24a/b in zebrafish embryos results in reductions in sarcomeric proteins, profound disorganization of the myofibril, looping defects, changes in heart rate, and reduced circulation (Maragh et al., 2011;
Poon et al., 2012). A second RBM protein, Rbm20, has been shown to regulate the splicing of transcripts encoding titin, a large sarcomeric protein (Guo et al., 2012; Li et al., 2013). Mutations in RBM20 have been linked with dilated cardiomyopathy in humans, consistent with an important role in regulating myofibril structure and/or function (Brauch et al., 2009; Li et al., 2010; Refaat et al., 2012). Loss of another RBM protein, Rbm15, in mice leads to a variety of embryonic defects, including cardiac abnormalities and heart failure (Raffel et al., 2009), though its role in heart development remains poorly defined.
RNA binding protein, fox-1 homolog (RBFOX) proteins are RRM-containing
proteins (Figure 4) that specifically bind to (U)GCAUG elements and regulate
alternative splicing (Kuroyanagi, 2009). In addition, RBFOX1 (also known as A2BP1)
57 has been shown to interact with ataxin-2, a protein of unknown function that
possesses two Sm domains also found in small nuclear ribonucleoproteins,
suggesting RBFOX1:ataxin-2 complexes may have additional roles in RNA
processing or transport (Shibata et al., 2000). RBFOX1 and RBFOX2 (also known as
RBM9) are highly expressed in the developing heart, skeletal muscle, and brain, and
(U)GCAUG motifs have been found to be enriched in transcripts that are
alternatively spliced in these tissues (Bland et al., 2010; Brudno et al., 2001; Das et
al., 2007; Gallagher et al., 2011; Jin et al., 2003; Sugnet et al., 2006; Zhang et al.,
2008a). The splicing activity of RBFOX proteins and splicing patterns of RBFOX
targets in the heart are highly conserved across species (Gallagher et al., 2011; Jin et al., 2003; Venables et al., 2012). Although knockdown of individual Rbfox proteins in zebrafish does not result in gross defects, rbfox1/rbfox2 double morphant embryos exhibit severe skeletal and cardiac muscle defects, including myofibrillar disorganization, disrupted swimming, and reduced cardiac function (Gallagher et al.,
2011). Expression of rbfox2 alone is sufficient to rescue wild type myocardial structure and function, suggesting that it can complement the functions of rbfox1 in cardiac myofibril development (Gallagher et al., 2011).
For proper assembly and maintenance of myofibrils, and for cardiomyocyte function, connections must be formed and maintained between apposing myofibrils, between myofibrils and the plasma membrane, and between neighboring cardiomyocytes. The RBP Fragile X mental retardation, autosomal homolog 1
(FXR1), has recently been shown to be involved in the proper regulation of these
connections (Whitman et al., 2011). FXR1, a homolog of FMR1, which is lost in
58 Fragile X Syndrome, contains two KH domains and an RGG box (Figure 4) (Burd and
Dreyfuss, 1994; Siomi et al., 1995). A role in transport and translation regulation of mRNA targets has been proposed for fxr1 during somitogenesis in Xenopus (Huot et
al., 2005). Fxr1-null mice have disorganized cardiac and skeletal muscle, and die
within a few hours of birth, likely due to cardiac and respiratory insufficiency
(Mientjes et al., 2004). Using these mice, Fxr1 was shown to regulate the expression
of Talin2, which is found at the costameres (i.e., myofibril-membrane anchors), and
Desmoplakin, which is found at the desmosomes (i.e., cell-cell junctions), in cardiac
muscle at the level of translational repression (Whitman et al., 2011). Dysregulation
of these proteins results in disruption of desmosome, costamere, and sarcomere
structures at the microscopic level, and in overt cardiomyopathy and muscular
dystrophy at the macroscopic level (Whitman et al., 2011). The expression and
function of Fxr1 in the developing heart and skeletal muscle is conserved in
zebrafish (Engels et al., 2004), where knockdown of fxr1 using antisense
morpholinos likewise results in abnormal myotome formation and severe
embryonic cardiomyopathy (Van't Padje et al., 2009).
3.3. Cardiac morphogenesis
Following formation of the primitive heart tube, the heart undergoes
extensive morphogenesis to transform from a simple tube into a multi-chambered
pump that is capable of directing pulmonary and somatic circulation. Left-right
asymmetry, established in the early embryo by a combination of asymmetrically
deposited signaling molecules and the asymmetric activation of transcription
59 programs, culminates in the heart in cardiac looping, in which right-handed bulging
and twisting of the heart tube gives rise to a C-shaped tube with the inflow and
outflow tracts both oriented rostrally (Brand, 2003). Differential proliferation
distinguishes the “outer curvature” from the “inner curvature” of the heart, and
contributes to subsequent expansion and definition of the chambers (Wagner and
Siddiqui, 2007). RBPs have been implicated in the regulation of morphogenesis of
the valves and septa that then divide the chambers, as well as morphogenesis of the
developing myocardium (Figure 3).
3.3.1. Endocardial cushion development
The looped heart tube is subdivided into a four-chambered heart through the
formation and remodeling of structures called endocardial cushions in the atrioventricular canal (AVC) and outflow tract (OFT) regions (DeLaughter et al.,
2011; Person et al., 2005). In the AVC and OFT, the endocardium is pushed away from the myocardium by the localized expansion of extracellular matrix. The cushions are cellularized by the invasion of mesenchymal cells produced from a subpopulation of endocardial cells via epithelial-to-mesenchymal transition (EMT).
Subsequent remodeling into the valves and septa involves condensation and differentiation of the cushion mesenchyme at post-EMT stages (Kirby, 2007). The
AVC cushions give rise to the mitral and tricuspid valves, as well as to the atrioventricular septum, and contribute to the atrial and ventricular septa (Kirby,
2007). The OFT cushions give rise to the aortic and pulmonary valves, and
60 transiently septate the OFT during early morphogenesis (Qayyum et al., 2001).
Dysregulation of EMT in the heart can lead to valve and septal defects (Person et al.,
2005). Although the tissue interactions, growth factors, signaling pathways, and transcription factors involved in regulating endocardial cushion EMT have been well studied (DeLaughter et al., 2011; Person et al., 2005), the roles of post- transcriptional RNA processing in regulating EMT and post-EMT remodeling are much less well understood.
A member of the muscleblind-like (MBNL) protein family, MBNL1, has been implicated in regulating endocardial cushion EMT (LeMasters et al., 2012; Vajda et al., 2009). MBNL proteins bind to RNA via two conserved pairs of zinc knuckle domains (Figure 3), and regulate pre-mRNA alternative splicing, RNA localization, and mRNA stability (Ho et al., 2004; Masuda et al., 2012; Wang et al., 2012). In the chicken embryo, MBNL1 expression is not detected in the linear heart tube prior to cardiac looping, but is detected in the looped heart prior to the formation of the endocardial cushions (LeMasters et al., 2012; Vajda et al., 2009). In the AVC and
OFT, MBNL1 is strongly expressed in the endocardium (LeMasters et al., 2012; Vajda et al., 2009). Knockdown of MBNL1 in chick AVC or OFT explants induces enhanced
EMT ex vivo, indicating that MBNL1 is a negative regulator of EMT in the endocardial cushions (LeMasters et al., 2012; Vajda et al., 2009). Active, secreted TGFβ3 levels are also elevated following MBNL1 knockdown (LeMasters et al., 2012). TGFβ proteins are important for inducing EMT (Arthur and Bamforth, 2011; Kruithof et al., 2012), suggesting MBNL1 may regulate EMT in part via modulation of EMT- inducing signals. MBNL1 preferentially binds to YGCY motifs in vitro and in vivo
61 (Goers et al., 2010; Wang et al., 2012), and these motifs are highly enriched in
known MBNL1-responsive transcripts (Gates et al., 2011; Ho et al., 2004; Wang et
al., 2012). Strikingly, a recent study investigating alternative splicing in breast
cancer cells found an enrichment of YGCY motifs near exons that exhibited
decreased inclusion following EMT (Shapiro et al., 2011). This suggests MBNL1 may
play a general role in regulating alternative splicing transitions during EMT.
A paralog of MBNL1, MBNL2, is also expressed in the embryonic heart
(Fardaei et al., 2002; Fernandes et al., 2007; Kanadia et al., 2003b; Liu et al., 2008),
but its temporal and spatial distribution within the developing heart, and whether it
plays a role in EMT, have not yet been investigated. Knockdown of mbnl2 in zebrafish does result in cardiac abnormalities, however, including dilation and disorganization of the myofibril, and mis-regulated alternative splicing of tnnt2, a known target of MBNL proteins in the myocardium (Machuca-Tzili et al., 2011).
Additional RBPs have also been implicated in regulating alternative splicing during EMT. Epithelial splicing regulatory proteins (ESRP) 1 and 2 are RRM-
containing RNA binding proteins that have been shown to regulate hundreds of
alternative splicing events during EMT (Dittmar et al., 2012; Warzecha et al., 2009a;
Warzecha et al., 2009b). Knockdown and over-expression studies have suggested
that changes in ESRP expression are determinative for EMT and its converse, MET
(Reinke et al., 2012; Shapiro et al., 2011; Warzecha et al., 2010), but in a recent
study neither Esrp1 nor Esrp2 were detected in the embryonic heart (Revil and
Jerome-Majewska, 2013). Motif analysis of intronic regions flanking EMT-regulated
cassette exons not only suggests common regulatory roles for MBNL and ESRP
62 proteins in EMT-specific alternative splicing, but also members of the polypyrimide tract binding protein (PTB), hnRNP, and RBFOX families (Shapiro et al., 2011). None of these have yet been interrogated for a role in endocardial cushion development.
3.3.2. Myocardial trabeculation and compaction
Myocardial cells are highly dynamic during cardiac morphogenesis,
undergoing epithelialization, proliferation, and compaction as trabeculae form and
then coalesce in the ventricular wall (Harvey and Rosenthal, 1999). The trabeculae
consist of protrusions into the lumen of the heart, and are made up of poorly
developed but strongly coupled cells, which contribute to the ventricular conduction
system (Moorman and Christoffels, 2003). The trabeculae also increase the surface
area of the thickening tissue, which is critical for oxygenation of the heart before the
coronary arteries form. Although the anatomical changes that occur during
trabeculation and compaction have been well described, the mechanisms that
control these processes are not well understood.
Cardiac helicase activated by MEF2 protein (CHAMP) is an RNA helicase
expressed specifically in the myocardium during prenatal and postnatal
development (Liu et al., 2001). CHAMP is a member of the RNA helicase superfamily
I, and shares several conserved motifs with helicases involved in DNA replication,
transcription, and RNA processing (Figure 4) (Liu et al., 2001). Embryonic
expression of CHAMP begins in the linear heart tube following the initiation of
MEF2C expression. During trabeculation, CHAMP is strongly expressed in the non-
63 proliferative trabecular cardiomyocytes, but not in the proliferative compact zone
(Liu et al., 2001). This regional localization, as well as spatiotemporal similarities
between the expression of CHAMP and both neurotrophin-3 and its receptor, Trk C,
which regulate cardiomyocyte proliferation (Lin et al., 2000), led to speculation that
CHAMP plays a role in repressing cardiomyocyte proliferation and growth during
myocardial morphogenesis (Liu et al., 2001). Although this has not been experimentally confirmed in vivo, over-expression of CHAMP in primary neonatal cardiomyocytes inhibits cellular hypertrophy and leads to the up-regulation of the cell cycle inhibitor p21 (Liu and Olson, 2002).
Interestingly, CHAMP is a variant of the testis-specific helicase, MOV10 like-1
(MOV10l1), generated by alternative promoter usage, using a start codon within
exon 14 (Liu et al., 2001). A second cardiac-specific helicase, cardiac-specific
isoform of Mov10l1 (Csm), is also generated by alternative promoter usage from the
MOV10l1 locus; its start codon lies within exon 16 (Ueyama et al., 2003). Despite
their similar origins, the two helicases have distinct effects. While Csm potentiates
cardiomyocyte hypertrophy induced by phenylephrine treatment (Ueyama et al.,
2003), CHAMP over-expression is able to block this hypertrophy (Liu et al., 2001).
The expression and role of Csm in embryonic heart development has not been
investigated.
3.4. Postnatal maturation of the heart
Although the four-chambered architecture of the heart is established during
embryogenesis, the rerouting of circulation to the lungs, switch from hyperplastic to
64 hypertrophic growth, and increase in workload on the heart after birth prompts
extensive molecular and cellular remodeling during early postnatal life. This
remodeling involves changes in the expression of growth factors, cell cycle
regulators, contractile and cytoskeletal proteins, and ion channels (Chen et al., 2004;
Harrell et al., 2007; MacLellan and Schneider, 2000; Siedner et al., 2003), and is not
limited to changes in transcription, but also includes changes in post-transcriptional
RNA processing.
3.4.1. CELF- and MBNL-mediated alternative splicing programs
Many cardiac transcripts undergo fetal-to-adult changes in alternative splicing (Kalsotra et al., 2008; Park et al., 2011). Several families of RBPs have been implicated in fetal-to-adult reprogramming of alternative splicing in the heart
(Figure 3). A study using splicing-sensitive microarrays identified fetal-to-adult splicing transitions in the developing mouse heart, many of which were conserved between mammals and birds (Kalsotra et al., 2008). Computational analyses
identified several motifs that were highly enriched near the developmentally-
regulated exons, including putative binding sites for hnRNP, PTB, STAR, RBFOX,
MBNL, and CUGBP, Elav-like family (CELF) proteins (Kalsotra et al., 2008).
Consistent with this, some of these RBPs were shown to be developmentally regulated. RBFOX1 transcript and protein levels are robustly up-regulated shortly after birth, whereas those of RBFOX2 decline slightly in the adult heart (Kalsotra et al., 2008). MBNL1 is up-regulated over the course of embryonic and postnatal heart
65 development (Kalsotra et al., 2008; Terenzi and Ladd, 2010). CELF1 and CELF2
exhibit higher protein levels in embryonic than adult heart, and are down-regulated
during early postnatal life (Kalsotra et al., 2008; Ladd et al., 2005a). Interestingly, the down-regulation of CELF proteins during heart development occurs without a change in CELF transcript levels, indicating that these RBPs are themselves post- transcriptionally regulated. Mechanisms include phosphorylation-driven changes in
CELF1 protein stability and miRNA-mediated repression of CELF1 and CELF2 translation (Kalsotra et al., 2010; Kuyumcu-Martinez et al., 2007).
Strikingly, over half of the fetal-to-adult splicing transitions identified by
Kalsotra and colleagues respond to over-expression of CELF1 or loss of MBNL1 in the hearts of genetically modified mice, suggesting these proteins are determinative for driving these developmental transitions (Kalsotra et al., 2008). A subset of these splicing events is regulated by both CELF1 and MBNL1 in an antagonistic manner.
Although a cardiac phenotype has not yet been described for Mbnl1-null mice
(Kanadia et al., 2003a), over-expression or repression of CELF proteins in the early postnatal myocardium leads to dysregulation of CELF-mediated alternative splicing and rapid onset of cardiac dysfunction in transgenic mice (Koshelev et al., 2010;
Ladd et al., 2005b; Terenzi et al., 2009). The juvenile onset of cardiomyopathy, as well as the spontaneous recovery of cardiac function following maturity in a line of mice with mild repression of CELF activity (Terenzi et al., 2009), further supports a role for CELF-mediated alternative splicing programs specifically in postnatal remodeling.
66 CELF and MBNL proteins are found in the cytoplasm as well as the nucleus in
the developing heart (Blech-Hermoni et al., 2013; Kalsotra et al., 2008; Ladd et al.,
2005a), suggesting that the effects of these RBPs in postnatal maturation are likely
not restricted to pre-mRNA alternative splicing. Although their cytoplasmic roles
have not been investigated in the heart, in developing skeletal muscle CELF1
regulates the translation of Mef2A and p21, key regulators of muscle-specific gene expression and growth arrest (Timchenko et al., 2004), and both CELF1 and MBNL1 have been shown to regulate the stability of a large number of muscle transcripts
(Masuda et al., 2012). Interestingly, CELF1 promotes decay of Mbnl1 transcripts and
MBNL1 promotes decay of Celf1 transcripts, suggesting that these factors mutually contribute to their reciprocal patterns of expression during myogenesis (Masuda et al., 2012).
3.4.2. Roles of the SR protein family in the maturing heart
Members of the serine/arginine-rich (SR) protein family have also been
implicated in fetal-to-adult cardiac reprogramming. There are twelve human SR
proteins, SRSF1-12, each characterized by the presence of one or two RRMs and a
carboxy-terminal RS domain enriched with arginine/serine dipeptides that
functions as a protein:protein interaction domain (Figure 3) (Twyffels et al., 2011).
SR proteins regulate splicing of both constitutive and alternative exons, mRNA
export, stability, and translation (Shepard and Hertel, 2009; Twyffels et al., 2011).
SR proteins also promote the processing of some miRNAs by facilitating cleavage by
67 Drosha (Wu et al., 2010a). The subcellular localization and activities of SR proteins are regulated in part through phosphorylation by SR protein-specific kinases (Zhou
and Fu, 2013). Depletion of SRSF1 (formerly known as ASF/SF2) induces apoptosis
in the DT40 chicken B-cell line (Li et al., 2005; Wang et al., 1996), and germline deletions of Srsf1, Srsf2 (formerly known as SC35), or Srsf3 (formerly known as
SRp20), result in early embryonic lethality (Jumaa et al., 1999; Wang et al., 2001; Xu
et al., 2005). Together, these studies have suggested essential, non-redundant roles
for multiple SR proteins in cell viability. Cardiac-specific knockouts of two of these
essential SR proteins, however, indicate that they have additional roles in regulating
contractile function in developing heart muscle.
Cardiac-specific ablation of Srsf1 or Srsf2 was accomplished by crossing mice with floxed alleles with the MLC-2v-Cre transgenic line (Ding et al., 2004; Xu et al.,
2005). Both cardiac-specific Srsf1- and Srsf2-null mice are healthy at birth, but develop early onset cardiomyopathy within the first four weeks of life (Ding et al.,
2004; Xu et al., 2005), corresponding to the period of postnatal remodeling.
Cardiomyocyte apoptosis was not observed in either model, but both displayed defects in excitation-contraction coupling (Ding et al., 2004; Xu et al., 2005). In cardiac-specific Srsf1-null mice this has been attributed at least in part to altered splicing of Ca2+/calmodulin-dependent kinase IId (CaMKIId), and transgenic over-
expression of the inappropriate CaMKIId splice form was sufficient to phenocopy
the defects in calcium handling in these mice (Xu et al., 2005).
SRSF10 (formerly known as SRp38) has also been implicated in regulating
calcium handling in the developing heart. Unlike SRSF1, SRSF10 is not essential for
68 viability in DT40 cells, although its loss does impair recovery from stress (Shin et al.,
2004). Germline ablation of Srsf10 does not result in the early embryonic lethality seen in other SR protein gene knockouts, but nonetheless few homozygous Srsf10- null fetuses reach full term (Feng et al., 2009). Most Srsf10-null embryos die by embryonic day E15.5 and exhibit multiple cardiac abnormalities, including septal defects, thinning of the myocardium, and altered intracellular calcium handling in embryonic cardiomyocytes (Feng et al., 2009). It was suggested that the calcium handling defects may be due to changes in the level and alternative splicing of triadin (Feng et al., 2009), though this is unlikely to contribute to the other developmental defects in Srsf10-null mice as triadin-null mice are viable with no obvious cardiac malformations (Shen et al., 2007). The earlier onset of cardiac dysfunction and presence of additional defects in Srsf10-null versus cardiac-specific
Srsf1- and Srsf2-null mice indicates that although multiple SR proteins are expressed in the heart, they play distinct roles in cardiac development.
3.5. Developmental dysregulation and disease
Dysregulation of developmental programs is often seen in disease states.
Disruption of RBP function during embryogenesis can disrupt proper formation of the heart, leading to congenital heart disease. Dysregulation of developmental RBP pathways in the adult heart, however, can also perturb cardiac function and contribute to acquired heart disease.
69 3.5.1. RNA binding proteins and congenital heart defects
Several RBPs have been linked with syndromes characterized by congenital
heart defects. Although a causal link has not been made in either case, CELF2 has
been proposed as a candidate gene for congenital heart defects associated with two
genetic disorders, partial monosomy 10p (Lichtner et al., 2002) and familial
arrhythmogenic right ventricular dysplasia (Li et al., 2001). A single case report has linked a partial deletion of the RBFOX1 gene with complex heart defects (Lale et al.,
2011), but further genetic evidence of RBFOX1 mutations causing developmental defects in humans is lacking.
Holt-Oram syndrome (HOS) is an autosomal dominant disorder characterized by upper limb abnormalities and a spectrum of cardiac birth defects, most typically septal and conduction defects (Mori and Bruneau, 2004). HOS is
caused by a variety of mutations in the gene encoding the transcription factor TBX5,
but the underlying mechanism of pathogenesis may not be limited to dysregulated
transcription. TBX5 has been shown to form an RNA-dependent complex with the
SR protein SRSF2, and affect constitutive and alternative splicing of reporter
minigenes in cells (Fan et al., 2009). Strikingly, a severe mutation in TBX5 (G80R)
associated with complete penetrance of cardiac defects strongly affects this splicing
activity, whereas a less severe mutation (R237Q) associated with incomplete
cardiac penetrance does not (Fan et al., 2009).
DiGeorge Syndrome (DGS) is caused in most cases by a deletion at the
genomic locus 22q11.2, and is characterized by a constellation of birth defects
attributable to disruption of neural crest cell development, including craniofacial
70 and cardiac defects (Keyte and Hutson, 2012). The transcription factor gene TBX1 is a leading candidate for DGS pathogenesis, but Tbx1 is not expressed in cardiac neural crest cells in mice (Garg et al., 2001; Vitelli et al., 2002). The DiGeorge critical region 8 (DGCR8) gene also lies within the 22q11.2 region, and encodes a double- stranded RBP essential for miRNA biogenesis (Seitz and Zamore, 2006). Genetic
inactivation of Dgcr8 specifically in neural crest cells in mice leads to cardiac
malformations typical of DGS, including persistent truncus arteriosis, aortic arch
malformations, and septal defects (Chapnik et al., 2012), supporting the idea that
loss of this RBP contributes to the cardiac defects in DGS patients.
Mutations in RBM10 have been implicated in talipes equinovirus, atrial septal
defect, Robin sequence (micrognathia, glossoptosis, and cleft palate), and
persistence of the left superior vena cava (TARP), a rare X-linked disorder with
severe congenital defects affecting multiple organs including the heart (Gripp et al.,
2011; Johnston et al., 2010). Although the molecular functions of RBM10 are not
well characterized, it was identified in a proteomic analysis of the human
spliceosome (Rappsilber et al., 2002), and has been shown to regulate the stability
of at least one mRNA (Mueller et al., 2009).
3.5.2. The recapitulation of fetal programs during adult heart disease
During heart disease, there is a partial reactivation of both transcriptional and post-transcriptional fetal gene expression programs. Many alternative splicing and alternative polyadenylation site choices found in the hypertrophic heart are
71 more similar to those found at fetal stages than in healthy adults, indicating that
patterns of fetal RNA processing are reestablished in response to pressure overload
(Ames et al., 2013; Park et al., 2011). These differences are likely mediated by RBPs
that become dysregulated during disease. For example, RBFOX1 is down-regulated while PTB is up-regulated during cardiac hypertrophy (Park et al., 2011).
Alterations in alternative splicing are associated with heart disease in both mice and humans (Ames et al., 2013; Kong et al., 2010; Park et al., 2011; Song et al., 2012).
Polymorphisms that affect alternative splicing of cardiac transcripts have also been
linked with susceptibility to myocardial infarction and cardiac hypertrophy
(Komamura et al., 2004; Mango et al., 2005).
Dysregulation of mRNA decay is another important driver of gene expression
changes during cardiovascular disease. The levels of AUF1 (also known as hnRNP
D), an RBP of the hnRNP family (Figure 3) that destabilizes transcripts via binding in
the 3’ untranslated region (3’ UTR) (Misquitta et al., 2001), are elevated in
cardiomyocytes in response to stress and in the hearts of human patients with heart
failure (Glaser et al., 2006; Pende et al., 1996). The increase in AUF1 is strongly
associated with a reduction in β-adrenergic receptor mRNA levels and impaired
calcium handling in heart failure (Misquitta et al., 2006). AUF1 is expressed in the
embryonic mouse heart (Gouble and Morello, 2000), but is normally undetectable in
the adult heart (Lu and Schneider, 2004), suggesting that this may also represent
the reactivation of a fetal decay program in adult heart disease.
Although the reemergence of fetal gene expression in the heart is generally
thought to be compensatory, the reiteration of fetal programs in adult tissues can
72 itself be pathogenic. In myotonic dystrophy (dystrophia myotonica, DM), the
expression of mutant RNAs containing expanded CUG or CCUG repeats leads to a
panoply of symptoms including electrocardiographic and echocardiographic
anomalies, skeletal muscle myotonia, muscle wasting and weakness, neurological
abnormalities, and endocrine dysfunction (Schoser and Timchenko, 2010). The
expression of expanded repeat-containing RNAs disrupts a number of RBPs in DM
cells, but pathogenesis is thought to be largely attributable to a gain of CELF1
function and loss of MBNL1 function that mirror embryonic expression patterns of
these factors (Schoser and Timchenko, 2010). The reiteration of fetal alternative
splicing patterns in adult DM tissues has been linked directly to patient symptoms
(Charlet-B. et al., 2002b; Savkur et al., 2001). Over-expression of CELF1 or deletion
of MBNL1 in mice is sufficient to reestablish fetal CELF/MBNL-mediated alternative
splicing patterns and mimic DM phenotypes (Ho et al., 2005; Kanadia et al., 2003a;
Koshelev et al., 2010; Timchenko et al., 2004; Ward et al., 2010). Conversely,
restoration of MBNL1 levels or repression of CELF activity rescues normal adult
alternative splicing patterns and reduces pathogenesis in a DM mouse model
(Berger and Ladd, 2012; Kanadia et al., 2006; Warf et al., 2009; Wheeler et al., 2009).
3.6. Concluding remarks
RNA binding proteins provide a robust and versatile mechanism for regulating gene expression. In eukaryotes, the regulation of alternative splicing of pre-mRNAs by RBPs underlies a great expansion in the proteome (Nilsen and
Graveley, 2010), allowing for the production of multiple gene products from the
73 majority of gene loci (Pan et al., 2008; Wang et al., 2008). RNA editing by RBPs can
not only expand the proteome (Maas, 2010; Rosenthal and Seeburg, 2012), but is
also important in the modification of non-coding RNAs (such as snoRNAs and
miRNAs) and in viral attenuation (Mallela and Nishikura, 2012). Transport, localization, and regulation of RNA stability by RBPs allow for direct control of the spatial and temporal profiles of gene products (Pratt and Mowry, 2013; Weis et al.,
2013), such as in the establishment of zygote polarity. Finally, regulation of translation by RBPs allows for the selective fine-tuning of protein production, and is particularly powerful in early zygotic events, during which maternal transcripts must be translated in the absence of a transcriptional apparatus (Lee and Schedl,
2006). All these mechanisms regulate gene expression without the need to transcribe new RNA, and thus make it possible to respond to external stimuli or developmental cues with remarkable speed and specificity.
This review describes the diverse RBP toolkit employed in the developing heart, which is involved in differentiation, morphogenesis, structure, and function.
In contrast to other tissues, such as the nervous system (Boutz et al., 2007; Gao and
Taylor, 2012; McKee et al., 2005; Okano and Darnell, 1997; Perrone-Bizzozero and
Bolognani, 2002; Yano et al., 2010), the identities and functions of RBPs in the developing heart have been pursued with significantly less vigor. While the importance of the heart cannot be disputed, its early appearance and small size in the embryo make the ability to investigate this developing organ technically difficult. Fortunately, the ability to investigate RNA processing (such as RNA editing, alternative splicing, or transcript occupancy by RBPs) using the small amounts of
74 tissue available from often microscopic embryos has recently become feasible thanks to the introduction of advanced methods such as laser-capture microdissection, high-throughput sequencing, and large-scale bioinformatic and computational analyses (Kishore et al., 2010; Wang et al., 2009).
A key challenge facing investigators as they adapt and apply these new tools is the identification of endogenous targets of the RBPs under study. For instance, while the splicing activity of regulators of alternative splicing has been investigated using artificial minigenes designed to mimic splicing substrates (Cooper, 2005), few bona fide targets have been identified for many of these regulators in vivo. At the same time, transcriptome analyses have indicated that the vast majority of genes (>
90%) are alternatively spliced (Pan et al., 2008; Wang et al., 2008), but have not linked these events to specific RBPs. Unlike DNA binding proteins, RBPs often do not have specific binding motifs. Instead, they bind with variable affinities to a range of sequences or secondary structures, with some preference for nucleotide content or morphology. This makes computational approaches to finding RBP binding sites challenging. Direct biochemical investigations of RBP binding have focused on identifying the highest affinity interactions, yet it is believed that most RBP:RNA interactions are not strong or are transient, and it is the relationships between RBPs and auxiliary factors that increase binding specificity (Burd and Dreyfuss, 1994;
Lunde et al., 2007). The role of weak or suboptimal binding in RNA processing remains to be elucidated (Pickrell et al., 2010).
With a large number of potential targets for each RBP, another important question to address is how these specific targets contribute to developmental
75 processes. In addition to directly affecting the expression of specific protein
isoforms involved in cell fate or morphogenesis, RBPs can exert control over key
regulatory proteins. For example, the activities of both transcription factors
(Belaguli et al., 1999) and splicing factors (Terenzi and Ladd, 2010) have been shown to be regulated in the heart by alternative splicing. Thus the identification and elucidation of RBP programs at key developmental stages may shed light on how signaling molecules, their receptors, and their downstream mediators are controlled at multiple levels. The burgeoning appreciation for the importance of
RBPs during normal development and in disease states, along with the rise of the technology necessary to properly interrogate their regulatory programs, will doubtless contribute important new insights into the formation and function of the developing heart. In time, RBPs and the transcripts they regulate during heart development may provide attractive targets for the design of treatments for congenital heart defects, cardiovascular disease, or cardiac tissue repair.
3.7. Acknowledgements
This work was supported by an NIH grant to A.N.L. (R01HL089376).
76 Chapter 4: Diversity and conservation of CELF1 and CELF2 RNA and
protein expression patterns during embryonic development
This chapter has been adapted from published work: Blech-Hermoni Y,
Stillwagon SJ, Ladd AN (2013) Diversity and conservation of CELF1 and CELF2 RNA
and protein expression patterns during embryonic development. Dev Dyn 242: 767-
777.
4.1. Introduction
CUG-BP, ELAV-like family (CELF) proteins are multifunctional RNA-binding
proteins (for a review, see (Dasgupta and Ladd, 2012)). Members of this family are evolutionarily conserved, and homologs have been identified in vertebrates and
invertebrates, as well as in plants (Brimacombe and Ladd, 2007). In this work, CELF
proteins will be referred to according to the most recent CELF nomenclature and,
where appropriate, other aliases will be included in parentheses. CELF proteins are
found in both the nucleus and the cytoplasm. In the nucleus, CELF proteins regulate
the alternative splicing of pre-mRNA targets, and CELF2 has been suggested to also
be involved in mRNA editing (Anant et al., 2001). In the cytoplasm, CELF1 and
CELF2 have been shown to regulate transcript stability and translation (Baldwin et
al., 2004; Iakova et al., 2004; Mukhopadhyay et al., 2003a; Timchenko et al., 2005).
The CELF proteins are divided based on sequence similarity into two
subfamilies, CELF1-2 and CELF3-6. Members of the two subfamilies differ in their
tissue distribution, with CELF1 and CELF2 being found in a broad array of tissues in
the adult (including heart, skeletal muscle, lung, liver, kidney, and the nervous
77 system) and CELF3-6 being found primarily in the nervous system (Ladd et al.,
2001; Ladd et al., 2004). The wide distribution of CELF1 and CELF2, in contrast to
the other members of the family, makes the expression and regulation of these
members of particular interest, especially in the context of the dynamic processes of
organogenesis.
While few studies have systematically compared the expression patterns of
these genes across species, developmental stages, or tissues, the available data
suggest that they are dynamically regulated during development. Celf2 (Napor)
transcript levels in the embryo increase from early to late embryonic development
in zebrafish and mouse (Choi et al., 1999; Choi et al., 2003; Levers et al., 2002). This
developmental regulation is also seen in Xenopus laevis, where total transcript levels
of all CELF members (Brunol1-5) increase during embryonic development, as
detected by RT-PCR on whole embryo RNA (Wu et al.).
The distribution of CELF transcripts has also been investigated in the
embryo, and suggests family member-specific regulation. In the zebrafish zygote, celf1 (brul) is found in the vegetal pole (Suzuki et al., 2000). By contrast, celf2
(napor) is found in the animal pole – as maternal transcripts – and its levels oscillate
sharply during early embryonic development (Choi et al., 2003). The limited
available data suggest that, at later stages, celf1 is strongly expressed in the lens
fiber cells of the developing zebrafish embryo, while celf2 is found in the nervous
system, somites, and retinal cells (Choi et al., 2003). In Xenopus laevis, celf1
(brunol2) of presumably maternal origin is found homogeneously in the early
embryo, although conflicting evidence suggests that these transcripts are localized
78 to the animal pole of the unfertilized egg and early-stage embryo (Wu et al.). At later stages, celf1 and celf2 (brunol3) transcripts are co-expressed in the Xenopus embryo, where their territories include the paraxial mesoderm, somites, neural structures, and the eye (Wu et al.). In chicken early embryonic development, CELF1 and CELF2 transcripts are found to be similarly expressed in the somites, nervous system, and heart muscle, while they localize to non-overlapping regions in the eye and endocardial cushions of the heart (Brimacombe and Ladd, 2007). In the developing mouse embryo, CELF1 and CELF2 transcripts have been detected in several brain regions, as well as in the eye (Choi et al., 1999; Diez-Roux et al., 2011; Levers et al.,
2002; Magdaleno et al., 2006; McKee et al., 2005).
In contrast to the patterns of transcript expression, the tissue distribution of the CELF1 and CELF2 proteins in the embryo has largely not been investigated.
Levels of both proteins have been shown to change from pre-natal to postnatal stages in several tissues including heart, skeletal muscle, liver, stomach, lung, and brain, but changes in protein levels during embryogenesis have not been studied
(Kalsotra et al., 2008; Ladd et al., 2001; Ladd et al., 2005a).
In this work, we compare CELF1 and CELF2 transcripts and proteins in different embryonic tissues, in order to characterize the levels and tissue, cellular, and subcellular distribution of these proteins in the developing embryo.
Furthermore, we compare the distribution of these CELF members in chicken and mouse embryonic development, in order to investigate the conservation of transcript and protein distribution in vertebrate embryogenesis. The expression patterns of CELF1 and CELF2 from this and previous studies are summarized in
79 Table 5. Our results demonstrate that CELF1 and CELF2 exhibit conserved, partially
overlapping, developmental stage-, tissue-, and subcellular compartment-specific
expression. These patterns of expression suggest that CELF1 and CELF2 may have
unique targets and roles in regions of the embryo in which the expression of one
family member is exclusive of the other.
4.2. Results and Discussion
4.2.1. CELF1 and CELF2 undergo dynamic post-transcriptional regulation during
early heart development in chicken
The heart first forms as a simple, linear tube (by stages 9-10, or roughly day
1-1.5). During cardiac morphogenesis, the primitive heart tube undergoes cardiac looping (stages 11-16, or roughly day 1.5-2.5) to orient the chambers and align the inflow and outflow tracts at the top of the heart. The looped heart then undergoes dramatic growth and rearrangement to form the morphologically familiar four- chambered structure (recognizable by stage 35, or day 8). Total protein levels of both CELF1 and CELF2 have previously been observed to drop between embryonic day 8 and adult (Kalsotra et al., 2008; Ladd et al., 2001; Ladd et al., 2005a). However, protein dynamics have not been investigated during the period of most active cardiac morphogenesis. To investigate the regulation of these proteins during this important period of heart development, the total protein levels of CELF1 and CELF2 were evaluated at eight stages between day 1.5 and day 14 of gestation. When total
80 CELF1 CELF2 Tissue Embryo Adult Ref.b Embryo Adult Ref.b Heart DB (H), ISH DB (H), WB 1-6 DB (H), ISH (M, DB (H); WB 1-6 (M, C), WB (M, C) C), WB (M, C) (M, C) (M, C), IF (M, C) Liver DB (H), ISH DB (H), WB 1-5 DB (H), ISH DB (H), WB 1-5 (C), WB (C), IF (M) (C), WB (C) (M) (M, C) Somites ISH (C) NA 5 ISH (C, Z) NA 5, 7, 8 Skeletal ISH (M, C, F), DB (H), WB 1-5 ISH (M, C), WB DB (H), WB 1-5, 9 muscle WB (M), IF (M) (M) (M) (M, C) Brain and DB (H), ISH DB (H), ISH 1-5, DB (H), ISH (M, DB (H), ISH 1-5, 7-9, nervous (M, C, F), WB (M), WB (M) 10, C, F, Z), RT- (M), WB (M) 11-13 system (M, C), IF (M, 11 PCR (M), WB C) (M) Eye ISH (M, C, F, ND 1, 5, ISH (M, C, F), ND 1, 5, 7, 9, Z), WB (C), IF 11, WB (C) 11 (M, C) 14 Lung DB (H), WB DB (H), WB 2, 3, 4 DB (H), WB DB (H), WB 2, 3, 4 (M) (M) (M) (M) Kidney DB (H), ISH DB (H), WB 1, 2, 3 DB (H) DB (H), WB 2, 3 (C) (M) (M) Spleen DB (H) DB (H), WB 2, 3 DB (H) DB (H), WB 2, 3, 4 (M) (M) G.I.c ISH (C), WB DB (H), WB 1-4 ISH (C), WB DB (H), WB 1-4 (M) (M) (M) (M) Repro., Md ND DB (H) 3 ND DB (H) 3 Repro., Fd ND DB (H), WB 2, 3 ND DB (H) 3 (M)
Table 5. Summary of CELF1 and CELF2 expression in embryonic and adult tissuesa aTranscript or protein expression was demonstrated in these tissues as indicated by Assay (Species): DB, RNA dot blot; ISH, in situ hybridization; RT-PCR, reverse transcription polymerase chain reaction; WB, western blot; IF, immunofluorescence; H, human; M, mouse; C, chicken; F, frog; Z, zebrafish; NA, not applicable; ND, not determined bReferences: (1) This study (Blech-Hermoni et al., 2013); (2) Ladd et al., 2001; (3) Ladd et al., 2004; (4) Ladd et al., 2005a; (5) Brimacombe and Ladd, 2007; (6) Kalsotra et al., 2008; (7) Choi et al., 2003; (8) Magdaleno et al., 2006; (9) Diez-Roux et al., 2011; (10) McKee et al., 2005; (11) Wu et al., 2010b; (12) Choi et al., 1999; (13) Levers et al., 2002; (14) Suzuki et al., 2000 cG.I., gastrointestinal tract dRepro., reproductive system; M, male; F, female
81
82 Figure 5. CELF1 and CELF2 expression in chicken heart during embryonic development. (A) Total CELF1 and CELF2 protein levels in whole heart rise from stage 10 to stages 23-26 (CELF1) and stages 26-29 (CELF2), and then drop, as determined by western blot. (B) Relative transcript levels of CELF1 and CELF2 do not vary significantly (p > 0.05; one-tailed t-test) in whole heart between stages 10 and 35, as determined by real-time RT-PCR (n = 3- 4). Error bars represent 95% Confidence Intervals. (C) Expression of CELF1 and CELF2 protein in heart nuclear and cytoplasmic fractions from stages 14, 23, and 35 as assessed by western blot. Ponceau S staining was used to show total protein integrity and loading. Bars by western blots indicate the relative position of the 50/52 kDa ladder marker for that blot. Blots shown are representative of three independent experiments.
83 protein samples from whole heart were analyzed by western blot, both CELF1 and
CELF2 appeared to be dynamically regulated. Both proteins were expressed at low
levels at stages 10 and 14, their expression then rose steeply — peaking between
stages 23 and 26 — followed by an acute decline (Figure 5A). Thus, the down- regulation reported at later embryonic and postnatal stages actually begins earlier than previously appreciated (Kalsotra et al., 2008; Ladd et al., 2001; Ladd et al.,
2005a). The specificity of the 3B1 antibody has been demonstrated previously using an array of different mouse tissues (Ladd et al., 2001) and transfected proteins
(Ladd et al., 2005a), and conditions were optimized for specificity of both antibodies in our hands.
While CELF1 and CELF2 protein levels decrease in mouse and chicken postnatally, their transcript levels remain unchanged, suggesting post- transcriptional regulation of CELF levels via changes in translation and/or protein stability (Kalsotra et al., 2008; Ladd et al., 2004; Ladd et al., 2005a). In order to investigate if CELF1 and CELF2 in the embryonic chicken heart are also regulated post-transcriptionally, we investigated the dynamics of expression of CELF1 and
CELF2 transcripts versus proteins. During the developmental window in which
CELF1 and CELF2 protein levels were dramatically rising and then falling, CELF1 and CELF2 transcript levels (measured by qRT-PCR) did not markedly change
(Figure 5B). These results are consistent with the regulation of these proteins via a post-transcriptional mechanism, as has been suggested postnatally.
CELF1 and CELF2 have been found in both nuclear and cytoplasmic fractions of mid-embryonic (E14) and early postnatal mouse heart, consistent with these
84 proteins carrying out both nuclear and cytoplasmic functions (Kalsotra et al., 2008;
Ladd et al., 2005a)(for review, see Dasgupta and Ladd, 2012). Since in previous studies the subcellular localization was determined at one embryonic stage, we investigated the dynamics of this localization over a broader developmental arc.
Western blots were performed on nuclear and cytoplasmic fractions of embryonic chicken hearts from several stages (Figure 5C), and probed for both
CELF1 and CELF2. Both nuclear and cytoplasmic CELF1 levels increased between stage 14 and 23, and then decreased between stage 23 and 35, as we have seen with total protein from whole heart. By contrast, while the nuclear CELF2 recapitulated the expression pattern in whole heart, the cytoplasmic CELF2 only decreased between stage 14 and 35. In this western blot assay, equal amounts of protein were loaded from both nuclear and cytoplasmic fractions (30 µg/sample, as described in
Chapter 2), which do not represent relative contribution to the total protein in the cell (e.g., the cytoplasm occupies a greater volume than the nucleus within the cell).
Therefore, a comparison of nuclear to cytoplasmic protein levels are not discussed here; rather, levels of nuclear protein from different stages or cytoplasmic protein from different stages are compared.
While the relative concentration of CELF1 in the nucleus was similar to
CELF1 in the cytoplasm, distinct nuclear (slower-migrating) and cytoplasmic
(faster-migrating) bands were identified (Figure 5C); a pattern that had not previously been reported. By contrast, CELF2 exhibited a doublet in cytoplasm and only a single band in the nucleus. Interestingly, this single nuclear band showed a developmental switch from a slow-migrating band at stage 14 to a faster-migrating
85 band at stages 23 and 35. These chicken cardiac CELF1 bands are indeed quite similar in relative mobility and their visualization is facilitated by separating the
protein samples over a longer period of time, while other CELF1 bands (such as seen
in eye; Figure 6) exhibit greater differences in relative migration.
Several sources of sequence variation have been reported for CELF1 and
CELF2, including variants in transcript sequence and post-translational
modifications of the proteins (partially curated in the UniProt database;
http://www.uniprot.org/uniprot/). There is evidence that both CELF1 and CELF2
are alternatively spliced, including one variant of CELF1 that codes for an extra four
amino acids (the “LYLQ variant”) and variants of CELF2 that would result in
peptides with extra amino acids (Choi et al., 1999; Ladd et al., 2001; Takahashi et al.,
2001). There is also evidence of alternative promoter usage at the CELF1 and CELF2
gene loci, giving rise to different primary transcripts in several tissues and cancer
cells yielding distinct protein isoforms with different amino termini ((Choi et al.,
1999)). Also, CELF1 and CELF2 contain several putative phosphorylation sites, and
phosphorylation of CELF1 at multiple sites has been associated with changes in
function and subcellular localization (Choi et al., 1999; Huichalaf et al., 2007; Li et
al., 2001; Ramalingam et al., 2008). Other post-translational modifications, such as
ubiquitylation, SUMOylation, and lipidation have not been investigated.
86
Figure 6. Expression of CELF1 and CELF2 proteins in a panel of chicken embryonic tissues. CELF1 and CELF2 protein expression in heart (he), liver (li), hindbrain (hb), and eye (ey) from stages 26 and 35 was assessed by western blot. Ponceau S staining was used to show total protein integrity and loading. Bars by western blots indicate the relative position of the 50/52 kDa ladder marker for that blot. Blots shown are representative of three independent experiments.
87 4.2.2. Different CELF1 and CELF2 isoforms are expressed in different tissues
Having found multiple isoforms of CELF1 and CELF2 in the developing heart,
we expanded our investigation to other tissues shown in the literature to express
these genes in the adult or embryo. Both CELF1 and CELF2 proteins have been
detected by western blot in a variety of adult tissues, including heart, skeletal muscle, liver, kidney, lung, and brain (Ladd et al., 2001; Ladd et al., 2005a). In this study, total protein from four tissues (heart, liver, hindbrain, and eye) at two developmental stages (stage 26, during which CELF protein levels peak in the heart and cardiac morphogenesis is underway, and stage 35, during which CELF protein levels in the heart are in decline and cardiac morphogenesis is largely complete) was compared by western blot analysis. In heart and liver samples, only a single
CELF1 band was resolved, whereas two bands were resolved in hindbrain and eye
(Figure 6). By contrast, a variety of bands were observed when the membranes were probed for CELF2, with little overlap between tissues. While the number of bands in each tissue did not change between stages, the relative abundance of protein in each band did appear to shift between stages 26 and 35 (Figure 6). In particular, the level of expression of the slow-migrating bands for both CELF1 and
CELF2 in eye and hindbrain decreased from stage 26 to stage 35.
An area of future investigation will be to distinguish the identity of these isoforms. Multiple mechanisms, including both differences in transcript sequences and post-translational modifications, may underlie CELF1 and CELF2 isoform diversity. Amplification of the full-length open reading frames of CELF1 and CELF2 from embryonic tissues by RT-PCR revealed several splice variants of each that
88 differ by three to eighteen nucleotides (data not shown). Although these differences
are too small by themselves to explain the differences observed by western blot
(encoding only one to six amino acids), they may affect the post-translational
modification of specific residues. Protein modifications may allow for dynamic
regulation of localization or activity of these proteins, and in these ways regulate the
class of substrates under CELF1 and CELF2 control during diverse conditions in
different tissues and developmental stages.
4.2.3. Regional distribution of CELF1 and CELF2 in the chicken embryo differ
between tissues
We have previously reported that, at earlier stages of development (stages
18 and 23), CELF1 and CELF2 transcripts are found in many of the same tissues
(such as in the nervous system, heart, and somites), and that their expression
territories overlap in some tissues (e.g., somites) but are non-overlapping in others
(e.g., the endocardial cushions of the heart)(Brimacombe and Ladd, 2007). In the
current study, we probed sagittal sections of chicken embryos from two more
advanced stages of embryonic development (stages 26 and 35) for CELF1 and CELF2 by in situ hybridization (Figure 7). Our data extend our previous findings, showing that CELF1 and CELF2 transcripts are co-expressed in some tissues (such as heart, liver, nervous system, and eye) at these stages as well, and that their territories overlap in some tissues (e.g., in the liver and in parts of the nervous system), while
89
Figure 7. CELF1 and CELF2 in situ hybridization in chicken embryos. Tissue distribution of CELF1 (A, C) and CELF2 (B, D) transcripts in sagittal sections of chicken embryos from stages 26 (A, B) and 35 (C, D; body only). Labels: he, heart; li, liver; br, brain; lu, lung; gi, gizzard; nt, neural tube; asterisks indicate dorsal root ganglia.
90 being spatially distinct in others (e.g., in the lens and retina)(Figures 8A-D). Our data go further to show that at stage 35, unlike at earlier stages, both CELF1 and
CELF2 are found in the gut; that CELF1 – but not CELF2 – is found in kidney at this stage; and that at both stage 26 and stage 35 CELF1 – but not CELF2 – is detectable at appreciable levels in the dorsal root ganglia.
The dichotomy of expression in the embryonic eye was also very striking at the protein level. When whole protein was extracted from isolated lenses and retinas of stage 26 or stage 35 embryos and analyzed by western blot, very sharp differences could be seen for both CELF1 and CELF2. Like their RNAs, CELF1 protein expression is higher in the lens, while CELF2 protein is more highly expressed in the retina. At both stages, a CELF1 band (possibly a doublet) was found in the lens, which was distinct from a CELF1 band found in retina, and the same was true for a lens-specific CELF2 band versus a retina-specific CELF2 doublet (Figure 8E).
Furthermore, our findings show that, while CELF2 transcripts are virtually undetectable in lens (while its relative levels are quite high in other tissues, especially brain), the protein is readily detectable by western blot (at levels equivalent or greater than those in brain), suggesting that either translation of
CELF2 is very efficient or the protein is particularly stable in the lens.
Our findings suggest that CELF1 and CELF2 are regulated both in terms of protein levels – in a manner that our data suggest is independent of transcript levels
– and in terms of their subcellular localization. Furthermore, while our findings by western blot analysis suggest the presence of multiple protein isoforms in different subcellular compartments, the protein samples represent multiple cell types from
91
92
Figure 8. CELF1 and CELF2 are differentially expressed in the embryonic chicken eye. CELF1 (A, C) and CELF2 (B, D) transcripts in transverse (A, B) and coronal (C, D) sections of stage 35 embryonic chicken heads (transverse and coronal planes of section for the embryo correspond to sagittal and transverse planes of section for the lens, respectively). Insets show magnified views of the lens in each panel. (E) CELF1 and CELF2 protein levels in lens and retina (ret.) of chicken embryos from stages 26 and 35, determined by western blot. Ponceau S staining was used to show total protein integrity and loading. Bars by western blots indicate the relative position of the 50/52 kDa ladder marker for that blot. Blots shown are representative of three independent experiments.
93
94 Figure 9. Tissue distribution of CELF1 protein in embryonic chicken. (A and B) CELF1 protein in sagittal sections of embryos from stages 26 (A) and 35 (B), determined by immunofluorescence. (C) High magnification (40X) images of CELF1 protein distribution in heart, liver, and hindbrain of an embryo from stage 35. (D) CELF1 protein in the whole eye of an embryo from stage 35; smaller panels show enlargements of lens and retina. Green, CELF1; blue, DAPI; each section is representative of two embryos. Labels: endo, endocardial cells; myo, myocardial cells; LEC, lens epithelial cells; LFC, lens fiber cells; TZ, transitional zone; GCL, glial cell layer; RPE, retinal pigment epithelium. Since retinal staining was variable and patchy, the high magnification view of the retina is from a different section than the one shown for the whole eye and the lens. This figure was modified from the published version for clarity.
95 each represented tissue. In order to visualize the relative levels, the subcellular distribution, as well as the sub-tissue distribution of CELF1 and CELF2 in the embryo, we employed indirect immunofluorescence (Figure 9). Despite attempts to optimize conditions (e.g., fixation, embedding, and antigen retrieval), immunofluorescence for CELF2 was unsuccessful for multiple antibodies (Fisher catalog number PA14130, Calbiochem catalog number 40-7500, Abcam catalog number ab111728, and 1H2 [Santa Cruz Biotechnologies])(not shown). While CELF1 protein was present in all transcript-positive tissues of stage 26 and stage 35 chicken embryos, its intracellular distribution in heart and skeletal muscles was strikingly different from its expression in other tissues. In particular, CELF1 was detected in both the cytoplasm and nucleus in cardiomyocytes, although it appeared to be expressed more strongly in the nucleus, while in other tissues analyzed its expression was overwhelmingly cytoplasmic (Figure 9C). Furthermore, CELF1 expression within several tissues was regionally specific. In the heart, CELF1 was exclusively detected in cardiomyocytes, while in the brain and retina it was predominantly detected in the luminal layers (Figure 9C). The strongest expression of CELF1 (protein and transcript) was found in the lens (Figure 9D), where the protein was predominantly detected in the fiber cells and in the cells of the transitional zone, whereas it was nearly undetectable in the lens epithelial cells.
Taking our protein data together (as detected by western blot and by immunofluorescence), our findings suggest that the different isoforms of CELF1 and
CELF2 in the eye may underlie more than just localization, since the protein is found in the cytoplasm of cells in both the retina and the lens. This territorial separation
96 between CELF1 and CELF2 in the eye has also been shown in zebrafish and Xenopus embryos, suggesting that there is high conservation of function for these proteins in the eye (Choi et al., 2003).
4.2.4. The distribution of CELF1 and CELF2 transcripts and CELF1 protein are largely conserved between chicken and mouse
To investigate the conservation of expression of CELF1 and CELF2, in situ hybridization was performed on whole sections of mouse embryos from days E11.5 and E14 of gestation for Celf1 and Celf2 transcripts (Figure 10). Our results show that the regional distribution of both transcripts recapitulates many of the patterns we find in the chicken embryo (e.g., high expression of Celf1 and Celf2 in the brain and in muscle), but not others (e.g., both Celf1 and Celf2 are strongly expressed in dorsal root ganglia)(Figure 10A-D compared with Figure 7A-D; note that the mouse embryo does not possess a gizzard). In addition, in the eye we find Celf1 to be highly expressed in the lens and Celf2 to be predominantly in the retina, as in chick
(Figures 10E-H). Finally, we also found both Celf1 and Celf2 expressed in a striated pattern in the mouse embryonic tongue (Figures 10E-H), while our chicken sections did not include the tongue.
When we probed similar sections of whole mouse embryos for CELF1 protein by immunofluorescence, CELF1 expression patterns largely recapitulated those found in the chicken embryo: CELF1 protein is highly expressed in the heart, other striated
97
98 Figure 10. Celf1 and Celf2 in situ hybridization in sagittal sections of mouse embryos. Celf1 (A, C) and Celf2 (B, D) transcript distribution at E11.5 (A, B) and E14 (C, D). (E-H) Celf1 (E, G) and Celf2 (F, H) transcripts in transverse (E, F) and coronal (G, H) sections of the heads of E14 embryos (transverse and coronal planes of section for the embryo correspond to sagittal and transverse planes of section for the eye, respectively). Tissue labels: he, heart; li, liver; br, brain; lu, lung; nt, neural tube; to, tongue; asterisks indicate dorsal root ganglia.
99
100 Figure 11. Tissue distribution of CELF1 protein in embryonic mouse. (A and B) CELF1 protein in sagittal sections of embryos from E11.5 (A) and E14 (B). (C) High magnification (40X) images of CELF1 protein in heart, liver, and hindbrain of an E14 embryo. Labels: epi, epicardial cells; myo, myocardial cells; endo, endocardial cells. (D) CELF1 protein in the whole eye of an E14 embryo; smaller panels show enlargements of lens and retina (green, CELF1; blue, DAPI). Labels: LEC, lens epithelial cells; LFC, lens fiber cells; NR, neural retina; RPE, retinal pigment epithelium.
101 muscle, liver, nervous system, and lens (Figures 11A-D). Of note was the expression
pattern of CELF1 in the heart. In embryos of both species, CELF1 protein in the heart
was restricted to the myocardial cell layer. In the chicken embryonic heart,
cytoplasmic CELF1 expression was markedly lower than nuclear expression, while
cytoplasmic expression of CELF1 in myocardial cells of the mouse embryonic heart
were in many cells similar to nuclear levels of the protein (Figure 11C compared with Figure 9C). Finally, while CELF1 was detected in the chicken embryonic retina
(albeit only in the cells lining the luminal aspect), no protein was detected in the retina of the mouse embryo; this is despite the fact that CELF1 expression in the lens was robustly detected in the embryos of both organisms (Figure 11D). This observation may represent differences in the development of the eye between these two species, as well as differences in antibody affinity for CELF1 in chicken versus mouse retina.
4.3. Concluding remarks
In this study, we investigated the expression of CELF1 and CELF2 transcripts and proteins in developing chicken and mouse embryos. We showed that CELF1 and
CELF2 protein levels are dynamically regulated during cardiogenesis, and that these dynamics are independent of transcript levels. We extended the previously reported developmental series for CELF1 and CELF2 transcript expression, showing that the
largely overlapping expression territories of CELF1 and CELF2 transcripts are maintained into later stages of development, but also showed that CELF1 and CELF2 are mutually exclusive in the kidney (in addition to the endocardial cushions, the lens, and the retina). We identified tissue-specific isoforms of both CELF1 and CELF2
102 in several tissues, and showed that some of these isoforms are restricted to the
nuclear or cytoplasmic subcellular compartment in the heart. Finally, we compared
CELF1 protein expression between the chicken embryo and the mouse embryo,
showing that the patterns of protein expression are largely conserved from birds to
mammals, with some subtle differences. This conservation in expression patterns
suggests that the function of these proteins is largely conserved during
development. The distribution of CELF1 protein in both the nucleus and the
cytoplasm of myocardial cells in both species suggests that it has important
activities in both compartments, while the subtle difference in its distribution
between these compartments may suggest an expansion of its cytoplasmic role in
cardiomyocytes of the developing mouse.
Important questions are raised in regions of overlapping expression: Are the
proteins working synergistically? Are they antagonizing each other? Are they
modified in a way that maintains a level of mechanistic or physical separation
between them? Better reagents and tools for detection of CELF2 protein in situ are desperately needed in order to address these issues. Lastly, our data suggest that
CELF1 may play predominantly different roles in different tissues. For instance, in liver, our data suggest that CELF1 plays predominantly cytoplasmic roles, which is in line with reports of CELF1 function in hepatocytes (i.e., translation control;
Fardaei et al., 2001; Michalowski et al., 1999; Timchenko et al., 2006; Timchenko et al., 1999). By contrast, previous reports on CELF1 function in the brain have focused on it splicing regulatory (i.e., nuclear) role, whereas our data suggest that the protein is predominantly expressed in the cytoplasm (Barron et al., 2010). In fact,
103 further dissection of CELF1 (and CELF2) protein expression in particular cell
populations in complex tissues such as brain and eye will be useful in identifying the
specific tissue components that are regulated by these proteins. Such information
would help to further uncover the processes that are globally regulated by these
proteins in all CELF1/2-expressing cells, as well as the tissue- or cell-type-specific
processes that are regulated by these proteins.
4.4. Acknowledgements
We would like to thank the laboratory of Dr. Oliver Wessely for technical
assistance with in situ hybridizations. We would also like to thank Dr. Judith A.
Drazba and Dr. John Peterson for assistance with the production of tiled images of embryo sections. This work was supported by a National Institutes of Health grant
to A.N.L. (1R01HL089376). Y.B-H. was supported by a National Institutes of Health
training grant (5T32GM008056-28).
104 Chapter 5: CUG-BP, Elav-like family member 1 (CELF1) regulates
MYH7B expression in cardiac muscle
This chapter has been submitted for publication: Blech-Hermoni Y*,
Dasgupta T*, Coram RJ, Stillwagon SJ, Ladd AN (2014) CUG-BP, Elav-like family member 1 (CELF1) regulates MYH7B expression in cardiac muscle.
* These authors contributed equally to this study
5.1. Introduction
Alternative splicing allows the production of multiple mRNA species from a
single gene. More than 90% of human genes are alternatively spliced (Pan et al.,
2008; Wang et al., 2008). The majority of alternative splicing events occur within
the coding region, and the different protein isoforms that arise often have distinct
functional consequences (Kalsotra and Cooper, 2011; Kelemen et al., 2013; Modrek
and Lee, 2002). Although less common, alternative splicing of 5’ and 3’ untranslated
regions (UTRs) is also important because it can insert or remove elements that
affect mRNA localization, stability, or translation. It has been estimated that as many
as 50% of disease-causing mutations affect splicing (Ward and Cooper, 2010). A
growing body of evidence indicates that the splicing of cardiac transcripts
profoundly affects heart function and pathogenesis. For example, cardiac troponin T
(TNNT2), a component of the contractile apparatus, has an alternative exon 5 that
increases the calcium sensitivity of the myofibril when included (Godt et al., 1993;
McAuliffe et al., 1990). A correlation has been found between mis-splicing of TNNT2
and congenital heart disease in human patients (Saba et al., 1996). Aberrant
105 regulation of TNNT2 splicing is also observed in myotonic dystrophy type 1 (DM1), and is thought to contribute to cardiac dysfunction in DM1 patients (Philips et al.,
1998). Alternative splicing of many cardiac transcripts is dysregulated in heart
disease in both mice and humans (Kim et al., 2014; Kong et al., 2010; Park et al.,
2011), and targeted disruption of splicing regulators in the heart causes
cardiomyopathy in mice (Ding et al., 2004; Koshelev et al., 2010; Ladd et al., 2005b;
Terenzi et al., 2009; Xu et al., 2005). Polymorphisms that affect alternative splicing
of cardiac transcripts have also been linked with susceptibility to myocardial
infarction and cardiac hypertrophy (Komamura et al., 2004; Mango et al., 2005).
The RNA binding protein CUG-BP, Elav-like family member 1 (CELF1)
regulates cell type- and developmental stage-specific alternative splicing of cardiac
transcripts (Kalsotra et al., 2008; Ladd et al., 2001; Ladd et al., 2005a). In the
developing heart, CELF1 is restricted to the myocardium, and is found
predominantly in the nucleus in cardiomyocytes (Blech-Hermoni et al., 2013;
Brimacombe and Ladd, 2007). CELF1 expression peaks in the embryonic heart
during cardiac morphogenesis, then drops during fetal through adult stages (Blech-
Hermoni et al., 2013; Kalsotra et al., 2008; Ladd et al., 2005a). It has been proposed
that the decline in CELF1 after birth drives fetal-to-adult transitions in alternative splicing of CELF targets during postnatal maturation of the heart (Giudice et al.,
2014; Kalsotra et al., 2008; Ladd et al., 2005a). Up-regulation of CELF1 in the adult heart contributes to the pathogenic reiteration of fetal splicing patterns in DM1 mouse models and human patients (Ladd et al., 2001; Philips et al., 1998;
Timchenko et al., 2001a; Wang et al., 2007). Over-expression of CELF1 in heart
106 muscle recapitulates many of these splicing defects and induces cardiomyopathy in
transgenic mice (Ho et al., 2005; Koshelev et al., 2010). Up-regulation of CELF1 and
changes in CELF-mediated splicing have also recently been implicated in diabetic
cardiomyopathy (Verma et al., 2013).
To identify targets of CELF1 in the developing heart, we performed cross-
linking immunoprecipitation (CLIP) for CELF1 from embryonic day 8 chicken
hearts. We identified a previously unannotated exon in MYH7B as a novel target of
CELF1-mediated alternative splicing regulation. Inclusion of this exon would lead to the insertion of a premature termination codon that would either produce a short, nonfunctional MYH7B peptide or destabilize MYH7B transcripts. Knockdown of
CELF1 in primary chicken embryonic cardiomyocytes leads to an increase in exon inclusion and decrease in MYH7B levels. MYH7B is also a target of the cardiac transcription factor MEF2A, and changes in Myh7b levels in the hearts of transgenic
mice with repressed CELF activity correlate with changes in Mef2a alternative
splicing. Taken together, these data support a model in which CELF1 regulates
MYH7B levels both directly through the alternative splicing of MYH7B transcripts
and indirectly through the alternative splicing of Mef2a transcripts.
107 5.2. Results
5.2.1. CELF1 binds predominantly to UG-rich intronic sequences in the embryonic
heart
To identify cardiac transcripts that directly bind to CELF1 in vivo, we performed CLIP using embryonic day 8 chicken hearts (Figure 12A). Of 564 different tags obtained, 26% (148) represented bacterial rRNA sequences (a common contaminant of CLIP), and 35% (197) could not be mapped with high confidence or mapped to more than one genomic locus. The remaining 39% (219
tags) were successfully mapped to the chicken genome using the BLAT search tool
on the University of California, Santa Cruz (UCSC) genome browser (Kent, 2002)
and/or BLAST search of the NCBI database (Altschul et al., 1997). We identified 170
tags within known genes, 13 within unknown genes (i.e., there is EST and/or cDNA
evidence supporting a transcript expressed from that genomic locus, but the gene is
unannotated and has no recognized homologs), and 36 within intergenic regions
(Table S1). It should be noted that due to the poor annotation of the chicken
genome, the majority of genes containing tags (>70%) were not annotated in the
UCSC genome browser; in these cases, homology and synteny with human and
mouse orthologs were used to identify the corresponding chicken genes.
Approximately 75% of the CELF1 CLIP tags within genes mapped to introns
(Figure 12B), consistent with the localization of CELF1 in the nucleus in embryonic
heart muscle cells (Blech-Hermoni et al., 2013), and its known role as a regulator of
pre-mRNA alternative splicing in the heart
108
Figure 12. Cross-linking immunoprecipitation (CLIP) of CELF1 from embryonic chicken heart. (A) CLIP was performed on embryonic day 8 chicken hearts using an anti-CELF1 antibody. Vertical red line indicates immunoprecipitated CELF1:RNA complexes following addition of an RNase inhibitor to block high levels of endogenous RNase activity (“low RNase” lanes); fully digested complexes (“high RNase”) run just above the size of immunoprecipitated CELF1 alone (western blot). (B) Distribution of CLIP tags within known genes.
109
110 Figure 13. CELF1 CLIP tags are enriched with UG motifs. (A) Incidence of dinucleotides within CELF1 CLIP tags that map to known genes. The dotted green line indicates the incidence expected if all dinucleotides were equally represented. (B) Incidence of hexanucleotides within CELF1 CLIP tags that map to known genes. The 20 most-frequent hexanucleotides are indicated in red. The dotted green line indicates the incidence expected if all hexanucleotides were equally represented. The sequences of the top hexanucleotides are shown, with UG dinucleotides within those motifs in red. (C) The distributions of tags containing different numbers of UG or CA dinucleotides are shown.
111 (Kalsotra et al., 2008; Ladd et al., 2001; Ladd et al., 2005a; Philips et al., 1998). In other cell types, CELF1 has also been shown to regulate mRNA adenylation status, stability, and translation in the cytoplasm, primarily through interactions with the 5’ or 3’ UTR. In an analysis of CELF1 targets identified by CLIP followed by high- throughput sequencing (HITS-CLIP) in a mouse myoblast cell line, C2C12, over 90% of exonic tags were found to map to 3’ UTRs (with a negligible number of tags mapping to 5’ UTRs)(Masuda et al., 2012). By contrast, we found that less than half of the exonic CLIP tags we identified were mapped to either a 5’ or 3’ UTR, with the majority of exonic tags mapping to internal coding exons (Figure 12B). It is important to note that our assay was not intended to be saturating with regard to
CELF1 bound tags, as only about 500 clones were sequenced. Furthermore, our assay represents binding of CELF1 to transcripts in embryonic heart tissue, rather than a cell line often used to study the differentiation of skeletal myoblasts. Finally, species differences (chicken versus mouse) may underlie some of the observed differences. Encouragingly, several tags were found within exons that are known to be alternatively spliced, raising the possibility that CELF1 regulates the alternative splicing of cardiac transcripts via both intronic and exonic binding sites.
An analysis of the frequency of dinucleotides within the CELF1 CLIP tags indicates that there is an enrichment of UG dinucleotides (Figure 13A).
Furthermore, all of the most frequently occurring hexamers are U- and G-rich and contain at least one UG dinucleotide (Figure 13B). This is consistent with known
CELF1 binding preferences, as CELF1 has been shown to bind with high affinity to
U/G-rich elements in vitro, in particular those containing UG dinucleotides
112 (Marquis et al., 2006; Mori et al., 2008b; Takahashi et al., 2000; Tsuda et al., 2009).
Likewise, MEME analysis of CLIP tags recently obtained from mouse C2C12 myoblasts identified a consensus binding motif for CELF1 containing repeated UGU elements (Masuda et al., 2012). UG dinucleotides are not non-specifically enriched by the CLIP procedure, as UGs are not enriched in CLIP tags from an unrelated RNA binding protein, Nova, that binds to YCAY motifs (Ule et al., 2003 and Figure 14).
The distribution of CELF1 CLIP tags containing different numbers of UG dinucleotides shows that the enrichment of UG motifs is widespread, and not the result of a subset of tags with an unusually high number of UGs (Figure 13C). In contrast, a large fraction of tags contain only one or no CA dinucleotides, which have the same G/C content, but appear with a frequency expected by random chance. The distribution of UG dinucleotides within exonic and intronic tags is similar, but amongst the exonic tags there are significantly more UG dinucleotides in tags that fall within UTRs than in those within coding regions (1-tailed t-test, P = 0.002).
Most well characterized splicing regulatory elements are found close to the splice site(s) they regulate. U/G-rich motifs have been found to be enriched in intronic regions proximal to cassette exons (i.e., within 200 nt) that are alternatively included in heart and skeletal muscle, and the enrichment of these putative CELF binding sites is highly conserved in sequence and position between human, mouse, chicken, and frog (Castle et al., 2008; Das et al., 2007). An RNA binding map recently generated for CELF1 on the basis of 24 alternative splicing events in skeletal muscle cells indicates that binding immediately upstream of an alternative exon is associated with exon skipping, whereas binding in the proximal downstream intron
113
Figure 14. Sequence enrichment in Nova CLIP tags differs from CELF1 CLIP tags. (A) Tags identified by CLIP analysis using anti-Nova antibodies are somewhat depleted for the UG dinucleotide, in contrast to CELF1-CLIP tags, while showing an enrichment for pyrimidine dinucleotides. (B) Analysis of hexamers revealed an enriched for sequences containing the YCAY motif, known to be the preferred binding motif for this protein.
114 is associated with inclusion (Masuda et al., 2012). We found that only a small fraction of intronic CELF1 CLIP tags fall within 500 nucleotides of a known splice site. Using the subset of 46 intronic tags that fall within genes annotated in the UCSC genome browser (Table S2), we found that less than 20% (9/46) are within 500 nt of at least one annotated splice site; only 15.2% are within 500 nt of the nearest upstream splice site, whereas 8.7% are within 500 nt of the nearest downstream splice site. Furthermore, closer examination of the introns containing these CELF1
CLIP tags revealed that most were unusually large (Table 6 and Table S2).
Although introns flanking alternative exons tend to be longer than those flanking constitutive exons, the average intron length in chick is less than 3,000 nt regardless of its position (Deutsch and Long, 1999; Kim et al., 2007). Only 39% of CELF1 CLIP tag-containing introns were less than 10,000 nt in length, however, and 13% were over 100,000 nt long (Table S2). To determine whether the intron length or position of intronic binding sites in our CLIP tag set is specific to the species, developmental stage, and/or tissue type, we performed similar analyses on published CELF1 CLIP tags from postnatal mouse hindbrain (Daughters et al., 2009).
As we saw in the embryonic chicken heart, a minority of intronic CELF1 CLIP tags from the postnatal mouse brain lie within 500 nt of the nearest upstream (10.1%) or downstream (6.5%) splice site, and the introns containing them are unusually large (Table 6). These observations are also not unique to CELF1. Ule and colleagues reported that almost two-thirds of intronic Nova CLIP tags from adult mouse brain were found within large (>10,000 nt) introns (Ule et al., 2003).
115
Intron length Distance from Distance from upstream splice site downstream splice site CELF1 CLIP tags, embryonic chicken heart (this study)a Mean 44,293+8,608 nt 21,063+5,271 nt 23,202+5,600 nt Median 23,569 nt 3,718 nt 6,804 nt Range 370 to 247,578 102 to 190,843 nt 103 to 184,011 nt nt CELF1 CLIP tags, postnatal mouse brain (Daughters et al., 2009) b Mean 77,984+11,456 32,441+4,669 nt 45,462+8,298 nt nt Median 17,712 nt 6,342 nt 7,244 nt Range 293 to 729,767 35 to 221,376 nt 15 to 670,693 nt nt Nova CLIP tags, adult mouse brain (Ule et al., 2003) b Mean 87,458+9,169 nt 33,905+4,081 nt 53,481+6,976 nt Median 32,058 nt 8,309 nt 12,524 nt Range 106 to 729,767 11 to 377,919 nt 8 to 727,004 nt nt aOnly intronic tags lying within chicken genes that are annotated in the UCSC genome browser were included in this analysis bAll CLIP tags reported by the authors to be intronic were included in this analysis
Table 6. Lengths of introns containing CLIP tags and relative position of tags within introns
116
Analysis of the intronic Nova CLIP tags from their study shows that very few are
found within 500 nt of an upstream (8.1%) or downstream (6.6%) splice site, and
confirms that tag-containing introns are on average very large (Table 6).
5.2.2. Alternative splicing of MYH7B inversely correlates with total transcript levels
and CELF1 expression
CELF-mediated alternative splicing has been implicated in regulating
contractile function in the heart (Dasgupta et al., 2013; Ladd et al., 2005b; Terenzi et al., 2009). In order to evaluate potential targets of CELF1-mediated alternative splicing regulation, several transcripts were chosen for further evaluation, based on the mapping of their corresponding CELF1-bound tag to intronic sequence. In
particular, we chose to evaluate candidates with a described role in the cytoskeleton
or in cardiac contraction or contractility. Since regulators of alternative splicing are
believed to bind proximally to the regulated event (such as alternative exons), we
performed RT-PCR using primers designed to amplify the nearest two exons on
either side of the tag locus. This approach would allow us to identify events in which
CELF1 binding to an intron results in inclusion or skipping of an upstream or
downstream proximal exon. Alternative splicing was evaluated in this manner in
chicken embryonic primary cardiomyocytes, following siRNA treatment (cells were
transfected 24 hpp and RNA was collected 72 hpt). Of several (~30) candidates that were tested in this way, and for which the RT-PCR reactions were successful, some
117
118 Figure 15. CELF1 regulates the inclusion of an unannotated exon in chicken MYH7B transcripts. (A) A CELF1 CLIP tag (green) maps to an intron within the coding region of MYH7B. RT-PCR using primers in upstream and downstream exons (indicated by half arrows) revealed the presence of a previously unrecognized exon (blue box) that is alternatively included in the embryonic heart. Red lollipop symbol represents stop codon predicted in the included exon. (B) Translation of the transcript sequence including this exon indicates that its inclusion would lead to the insertion of an in-frame stop codon close to the N-terminal end of the protein. (C) Western blots were performed on total protein samples collected from primary embryonic cardiomyocytes mock transfected or transfected with siRNAs against CELF1 (si1 and si2) or a control siRNA (siCont). Representative blots from one of three independent transfections are shown. (D) The extent of inclusion of the novel MYH7B alternative exon was determined by semi-quantitative RT-PCR in primary embryonic cardiomyocytes grown and transfected in parallel to those used to harvest protein. Data represent mean values from three independent transfections. (E) Total MYH7B transcript levels in primary cardiomyocytes transfected with control (siCont) or anti-CELF1 (si2) siRNA were compared to mock-transfected controls by qRT-PCR. An asterisk indicates p ≤ 0.05 compared to mock. (F) Total Myh7b transcript levels were determined in the hearts of 24 week-old wild type and MHC-CELF∆-10 females by qRT-PCR. In this line, penetrance of cardiomyopathy is complete in females, while only about 50% of males (“affected”) exhibit cardiomyopathy. (G) Total Myh7b transcript levels were determined in the hearts of 24 week-old wild type, “unaffected” and “affected” transgenic MHC-CELF∆-10 males by qRT-PCR. (H) Total Myh7b transcript levels were determined in pectoral muscles of 24 week-old wild type and Myo- CELF∆-370 females by qRT-PCR. All mouse qRT-PCR data represent mean values from three different individuals. An asterisk indicates P ≤ 0.05 compared to wild type. Mouse lines are described in the text.
119 produced only one amplicon following all treatments, suggesting the absence of
alternative splicing regulation of the amplified exons in these cells. The remaining
candidates produced two amplicons, of sizes consistent with the inclusion/skipping
of one of the flanking exons. For a handful of candidates in this category, the proportions of the two variants changed following CELF1 knockdown, although the
sequences of most of these variants could not be confirmed. For most of the
candidates for which two variants were detected, however, the ratio of included
variant to skipped variant did not change following siRNA-mediated knockdown of
CELF1, suggesting that these events were unlikely to be regulated by CELF1 in these
cells. A product of an unexpected size was identified for one CELF1 CLIP tag, which
mapped to the first annotated intron of the chick MYH7B gene (Figure 15A). MYH7B
encodes a myosin heavy chain protein expressed in the embryonic heart, which is
incorporated into the thick filaments of the myofibril (Warkman et al., 2012). To
determine whether CELF1 binding could regulate the inclusion of the adjacent
downstream exon, semi-quantitative RT-PCR was performed using primers in the
flanking exons on total RNA from primary chicken embryonic cardiomyocytes
following CELF1 knockdown. Alternative splicing of the annotated upstream exon
could not be assessed by RT-PCR, as any sequences upstream of this exon in chick
transcripts remain unknown. Surprisingly, although the downstream exon was
constitutively included, a second, unexpected PCR product of slightly higher
molecular weight was identified. Gel isolation and sequencing of the PCR products
and alignment against the chick MYH7B genomic sequence revealed the variable
inclusion of a previously unannotated 84-nt exon located just 27 nt upstream of the
120 CLIP tag (Figure 15A). Inclusion of this exon would be predicted to lead to the insertion of an in-frame stop codon (Figure 15B). The level of inclusion of this exon inversely correlates with CELF1 protein levels in chick primary embryonic cardiomyocytes (Figure 15C,D), suggesting CELF1 inhibits exon inclusion. This is in contrast to the RNA binding map generated for CELF1 in skeletal muscle, where negative regulation of exon inclusion by CELF1 is associated with binding in the upstream intron, and enhanced inclusion is associated with binding downstream
(Masuda et al., 2012).
The stop codon encoded by the novel alternative exon is close to the N- terminus of the protein, preceding the first known functional domain. Its inclusion, therefore, could lead to the production of a truncated, nonfunctional myosin peptide. Alternatively, inclusion of this exon could lead to destabilization of the transcript via the nonsense mediated decay (NMD) pathway, which recognizes transcripts containing premature termination codons and targets them for destruction (Palacios, 2013). It should be noted that if this is the case, the relative fraction of MYH7B transcripts including this exon may be higher than shown in
Figure 15D, as unstable transcripts would be under-represented in measurements of steady state levels. Total MYH7B transcript levels are indeed lower in primary cardiomyocytes when exon inclusion is higher (Figure 15E), consistent with NMD.
121 5.2.3. CELF-mediated alternative splicing of Mef2a correlates with Myh7b levels in
transgenic mice
Using the conservation track of the UCSC genome browser, the region of the
MYH7B gene containing the novel exon and CLIP tag shows poor conservation in mammalian species. By RT-PCR we found no evidence of an alternative exon in the same relative position in the mouse Myh7b gene (data not shown), but alternative splicing of another CELF target may provide another means of regulating Myh7b levels. The Myh7b promoter contains highly conserved myocyte enhancer factor 2
(MEF2) binding sites, which were shown to be required for cardiomyocyte-specific
expression of an Myh7b reporter (Warkman et al., 2012). We previously demonstrated that Mef2a exon 16 inclusion is decreased in the hearts of MHC-
CELF∆ transgenic mice, which express a nuclear dominant negative CELF protein in
the myocardium (Dasgupta et al., 2013; Ladd et al., 2005b; Terenzi et al., 2009).
Skipping of exon 16 reduces the transactivation activity of MEF2A (Zhu et al., 2005).
In MHC-CELF∆-10 (high-expressing) transgenic males, which exhibit approximately
50% penetrance, decreased Mef2a exon 16 inclusion correlates with the
development of overt cardiomyopathy. “Affected” MHC-CELF∆-10 males (defined as
having a heart size greater than two standard deviations above the mean size of
their wild type counterparts) exhibited greater decreases in Mef2a exon 16
inclusion than “unaffected” MHC-CELF∆-10 males (defined as having a heart size
within one standard deviation of the mean size of wild type males) (Dasgupta et al.,
2013). To determine whether Myh7b levels correlate with Mef2a alternative splicing
in MHC-CELF∆ mice, Myh7b levels were determined by real time RT-PCR in the
122 hearts of MHC-CELF∆-10 females (which exhibit 100% penetrance), and both
“affected” and “unaffected” MHC-CELF∆-10 males. Indeed, Myh7b levels were
reduced in the MHC-CELF∆ mice (Figure 14F,G), and the extent of this decrease
correlated with the extent of Mef2a exon 16 skipping (Dasgupta et al., 2013).
MEF2A regulates muscle gene transcription in skeletal as well as cardiac
muscle (Edmondson et al., 1994). Myo-CELF∆-370 (high-expressing) mice express
the same nuclear dominant negative protein in skeletal muscle that MHC-
CELF∆ mice express in the heart (Berger et al., 2011). Surprisingly, however, we
found no significant effect on Mef2a exon 16 inclusion in pectoral muscles from
Myo-CELF∆ mice despite changes in splicing of other CELF targets (Berger et al.,
2011). Total Mef2a levels also do not differ between wild type and Myo-CELF∆
muscles (data not shown). Consistent with a lack of perturbation of MEF2A activity,
Myh7b transcript levels do not differ between the pectoral muscles of Myo-CELF∆
and wild type mice (Figure 15H).
5.3. Discussion
In this study, we performed CLIP for CELF1 on mid-gestation embryonic
chicken hearts. CELF1 CLIP had previously been reported from postnatal mouse
brain (Daughters et al., 2009), and CLIP-seq from mouse C2C12 myoblasts (Masuda
et al., 2012). Consistent with these reports and in vitro binding studies (Marquis et
al., 2006; Mori et al., 2008b; Takahashi et al., 2000; Tsuda et al., 2009), we found that
CELF1 binds to sequences enriched in UG dinucleotide motifs. Although the majority
of CELF1 CLIP tags fall within introns, which would be consistent with CELF1’s
123 localization in the nucleus in myocardial cells (Blech-Hermoni et al., 2013) and its
role as a regulator of pre-mRNA alternative splicing in the heart (Kalsotra et al.,
2008; Ladd et al., 2001; Ladd et al., 2005a), binding alone does not necessarily
denote regulation. It was recently reported that only about 12% of exons predicted
to undergo muscleblind-like 2 (MBNL2)-dependent splicing regulation based on
MBNL2 CLIP tag clusters exhibited altered splicing in microarrays using exon
junction probes or RNA-seq data from Mbnl2-null mice (Zhang et al., 2013). The
predominance of CELF1 binding in large introns far from known splice sites may suggest that a large percentage of intronic CELF1 binding is in fact nonfunctional.
Given their propensity to bind to short, degenerate motifs, it is not surprising that
binding of CELF1 and other RNA binding proteins can be found within very large
introns, which would be likely to harbor such motifs merely by chance. The UV
cross-linking used in the CLIP method has the potential to capture transient, low
affinity interactions as well as stable, high affinity interactions. That is not to say
that such binding would necessarily be without consequences within the cell. A
large number of nonfunctional binding sites could act as a molecular sponge,
“soaking up” and limiting the amount of an RNA binding protein available to interact
with its targets. Both coding and non-coding RNAs have been proposed to act as
molecular sponges for microRNAs, reducing the available pool that can bind and
regulate their target mRNAs (Kartha and Subramanian, 2014). Alternatively, binding
within these large introns could function to repress the use of pseudo splice sites,
regulate as yet unannotated exons (such as the novel exon we identified in MYH7B),
or act on true splice sites at a distance. Although most known splicing regulation
124 occurs via binding to proximal elements (Cooper, 2005), recent studies have shown
that distal intronic elements can dictate splice site choice in some cases (Lovci et al.,
2013; Parra et al., 2012).
We identified a novel CELF1-regulated exon in the first intron of the chicken
MYH7B gene that had a proximal CELF1 CLIP tag located 27 nt downstream.
Inclusion of this exon is predicted to target MYH7B transcripts for NMD, and indeed a decline in MYH7B levels is observed when inclusion of this exon is stimulated by
CELF1 knockdown in primary embryonic cardiomyocytes. Another group reported that skipping of a downstream exon, exon 7, leads to the introduction of a premature termination codon and down-regulation of Myh7b transcripts in mouse cells (Bell et al., 2010). It is not known whether exon 7 skipping occurs in chicken
MYH7B transcripts, but it is possible that the different species employ a similar
mechanism yet use different exons, or that chicken MYH7B possesses an additional
layer of splicing-mediated regulation not observed in mammals. An advantage of
splicing-based NMD as a mechanism for regulating MYH7B levels is that it enables
the cell to separate steady state levels of MYH7B protein from transcription of the
MYH7B gene, which also hosts a microRNA, miR-499, within one of its introns (Bell
et al., 2010). MYH7B has been shown to incorporate into the sarcomere, but how
MYH7B is distinct from other myosin heavy chain proteins in the contractile
apparatus is unknown. MYH7B expression is up-regulated during induced cardiac
hypertrophy in mice (Warkman et al., 2012), and a loss-of-function mutation in the
MYH7B gene has been proposed to underlie congenital myopathy with left
ventricular non-compact cardiomyopathy when combined with a mutation in the
125 integrin alpha 7 (ITGA7) gene in human patients (Esposito et al., 2013). These data suggest that expressing the right level of MYH7B in heart muscle is important for healthy contractile function.
Transcription of Myh7b is MEF2-dependent (Warkman et al., 2012), and here we show that Myh7b levels correlate with inclusion of an exon that is required for the transactivation activity of MEF2A in mouse striated muscle. The inclusion of
MEF2A exon 16 was not affected in chick primary embryonic cardiomyocytes following robust knockdown of CELF1 (si2; data not shown), suggesting the drop in
MYH7B levels in these cells is due to splicing of MYH7B, not MEF2A. The lack of
change in MEF2A exon 16 inclusion following CELF1 knockdown could be due to
compensation by CELF2, which is also expressed in the myocardium (Blech-
Hermoni et al., 2013; Brimacombe and Ladd, 2007; Ladd et al., 2001; Ladd et al.,
2005b), binds to similar U/G-rich motifs (Faustino and Cooper, 2005; Marquis et al.,
2006), and regulates splicing in a manner similar to CELF1 (Barron et al., 2010;
Gromak et al., 2003; Ladd et al., 2001; Ladd et al., 2004). In contrast, in MHC-CELF∆
mice the expression of a dominant negative CELF protein inhibits the activities of
both CELF1 and CELF2. Alternatively, differential splicing of MYH7B and MEF2A
transcripts could reflect different regulatory mechanisms employed in chicken versus mouse. Some differences in MYH7B expression patterns have been reported between species (Warkman et al., 2012).
It is unlikely that down-regulation of Myh7b is the full extent of the consequences of reduced Mef2a exon 16 inclusion, as MEF2A regulates the
transcription of many cardiac genes (Ewen et al., 2011; Naya et al., 2002). Among
126 these is four and a half LIM domains 2 (Fhl2). MEF2A binds to the Fhl2 promoter and activates its transcription, and Fhl2 levels are reduced in Mef2a-knockout mice
(Ewen et al., 2011). FHL2 interacts with and inhibits another cardiac transcription factor, serum response factor (SRF) (Philippar et al., 2004). SRF regulates the
transcription of a large number of cardiac genes involved in contractile function,
and plays roles in normal development, pathogenesis, and aging in the heart (Balza
and Misra, 2006; Nelson et al., 2005; Zhang et al., 2003). We previously demonstrated that Fhl2 transcript and protein levels are reduced in MHC-CELF∆ mice, consistent with a decrease in MEF2A activity, and this is accompanied by an increase in SRF targets (Dasgupta et al., 2013).
CELF1 regulates alternative splicing in skeletal as well as cardiac muscle
(Charlet-B. et al., 2002b; Savkur et al., 2001; Ward et al., 2010). Although repression of CELF activity in heart leads to cardiomyopathy and severe cardiac dysfunction in
MHC-CELF∆ mice (Ladd et al., 2005b; Terenzi et al., 2009), Myo-CELF∆ transgenic mice that express the same dominant negative protein under a skeletal muscle- specific promoter have a much milder phenotype despite similar levels of transgene expression (Berger et al., 2011). Some of this difference may be explained by differences in the composition of the CELF program, as some CELF targets are expressed only in heart (e.g., TNNT2), while others are expressed only in skeletal muscle (e.g., ClC1) (Charlet-B. et al., 2002b; Ladd et al., 2001; Philips et al., 1998).
Mef2a, however, is expressed in both heart and skeletal muscle (Edmondson et al.,
1994), yet in vivo its alternative splicing responds to loss of CELF activity in MHC-
CELF∆ hearts but not Myo-CELF∆ skeletal muscle (Berger et al., 2011; Dasgupta et
127 al., 2013; Ladd et al., 2005b; Terenzi et al., 2009). Consistent with Mef2a exon 16 splicing, Myh7b and Fhl2 levels also remain normal in Myo-CELF∆ skeletal muscle
(Figure 15H and data not shown). This indicates that whether CELF activity is determinative for exon inclusion for some targets depends on the tissue. The reason for this is unknown, but may reflect tissue-specific differences in other splicing factors that participate in their regulation. Alternative splicing outcomes are the result of combinatorial, often complex, control involving multiple RNA binding proteins that recognize distinct or overlapping cis elements in the pre-mRNA
(Barash et al., 2010; Hertel, 2008). The lack of perturbation of MEF2A, and subsequent FHL2-mediated inhibition of SRF, in Myo-CELF∆ muscle may explain the mildness of the skeletal muscle phenotype in comparison to the severely affected
MHC-CELF∆ heart muscle.
5.4. Concluding remarks
Using CLIP, we found that CELF1 binds to UG-rich motifs in cardiac transcripts. Many of these binding sites fall within large introns far from known splice sites. We also identified a novel target of CELF1-mediated regulation in heart muscle, MYH7B. We provide evidence that suggests CELF-mediated alternative splicing regulates MYH7B transcript levels by two separate mechanisms: directly via regulation of a previously unidentified MYH7B exon, and indirectly via regulation of an alternative exon in transcripts encoding the transcription factor MEF2A (Figure
16). Thus, CELF1 can modulate cardiac gene expression at both transcriptional and post-transcriptional levels.
128
Figure 16. Model for direct and indirect regulation of MYH7B expression via CELF-mediated alternative splicing. In chick primary embryonic cardiomyocytes, loss of CELF1 increases inclusion of a novel MYH7B alternative exon containing a premature stop codon, which would be predicted to destabilize MYH7B transcripts via nonsense-mediated decay or lead to the expression of a short, nonfunctional MYH7B peptide. In the mouse heart, repression of CELF activity stimulates skipping of Mef2a exon 16, which is known to reduce MEF2A’s transcriptional activity. This would also lead to a decline in MEF2A target genes, including Myh7b. Thus, CELF-mediated alternative splicing has the potential to regulate the expression of MYH7B both directly and indirectly, by modulating the splicing of MYH7B transcripts or the upstream transcription factor that regulates their production.
129 5.5. Acknowledgements
We thank Dara Berger, Natalie Vajda, and Kathryn LeMasters for assistance
with maintaining the transgenic mice and collecting mouse tissue samples. This
work was supported by a National Institutes of Health grant to A.N.L.
(1R01HL089376). Y.B-H. was supported in part by a National Institutes of Health training grant (5T32GM008056).
130 Chapter 6: CUG-BP, Elav-like family member 1 (CELF1) regulates
multiple aspects of myocardial development
This chapter is in preparation for publication: Blech-Hermoni Y, Wessely O, Ladd
AN. CUG-BP, Elav-like family member 1 (CELF1) regulates multiple aspects of myocardial development.
6.1. Introduction
CUG-BP, Elav-like family member 1 (CELF1) is a versatile RNA binding protein found in a variety of embryonic and adult tissues (Dasgupta and Ladd,
2012). CELF1 can regulate target transcripts by regulating their alternative splicing, stability, and translation. In the heart, CELF1 is expressed exclusively in the contractile cells of the myocardium, where its expression peaks during cardiac morphogenesis (Chapter 4; Blech-Hermoni et al., 2013). Knockdown of CELF1 in cultured chicken primary embryonic cardiomyocytes results in altered splicing of
MEF2A, a transcription factor important for muscle-specific expression programs and down-regulation of MYH7B, a myosin heavy chain incorporated into myofibrils and found in the embryonic heart (Chapter 5). The alternative splicing of another component of the myofibril, Cardiac Troponin T (TNNT2) can also be regulated by
CELF1 (Philips et al., 1998). In the human muscular dystrophy, Myotonic Dystrophy
Type 1, CELF1 is aberrantly upregulated and hyperactivated, resulting in defects of skeletal muscle differentiation, contractile dysfunction, and inappropriate splicing of target transcripts. Repression of CELF activity in the mouse heart post-natally
131 results in profound dilated cardiomyopathy, contractile defects, cardiac
hypertrophy, and cardiac fibrosis (Ladd et al., 2005b), but its role in the developing
myocardium has not been studied.
Beating of striated muscle cells is achieved by the contraction of myofibrils.
These intracellular fibers consist of linearly arranged sarcomeres, the basic units of
the contractile apparatus. The sarcomere is comprised of hundreds of proteins that work together to integrate intracellular release of calcium ions (Ca2+) from intracellular stores in the sarcoplasmic reticulum into physical force generation.
Structurally, each sarcomere consists of arrays of interdigitated thin Actin filaments
and thick Myosin filaments. The thin filaments are anchored within the structures
referred to as Z-discs, while the thick filaments are anchored within the M-lines.
While the thin and thick filaments are in close apposition, direct contact between
the Myosin heads and the Actin filaments in inhibited by a thin filament-associated
protein complex, the trimeric ‘Troponin Complex’, composed of a calcium binding
subunit (Troponin C), a Tropomyosin-binding subunit (Troponin T), and an
inhibitory subunit (Troponin I) that allows Tropomyosin to binds the complex to the
thin filament while at rest. Upon Ca2+ release into the cytoplasm, Troponin C binding to the cations mediates translocation of the Troponin I subunit to Actin binding by the Myosin heads. The Myosin then translocates along the actin filament in cyclical fashion mediated by ATP hydrolysis. This translocation bring the Z-discs closer together and translates into axial shortening of the myofibril. Myofibrils are anchored to the cell membranes laterally (at the costameres) and terminally (at the intercalated discs), allowing myofibrillar shortening to result in cellular contraction.
132
CELF proteins have been shown to regulate alternative splicing of transcripts
encoding contractile proteins, such as Alpha Actinin 1 (Gromak et al., 2003; Suzuki et al., 2002) and Cardiac Troponin T (Goo and Cooper, 2009; Ladd et al., 2001), as well as proteins that mediate interactions between muscle cells and the underlying substrate (Integrin β1; Itgb1; Ladd et al., 2005b). Furthermore, CELF1 has been implicated in muscle structure and function in both skeletal (Milne and Hodgkin,
1999; Timchenko et al., 2004) and cardiac (Ladd et al., 2005b) muscle. We hypothesized that CELF1 is involved in myofibrillar structure in cardiomyocytes of the developing heart and that disruption of CELF1 expression will result in myofibrillar defects. In order to test this hypothesis, we employed cellular approaches to study the effects of CELF1 depletion in cultured primary embryonic cardiomyocytes, and also evaluated the effects of CELF1 depletion in the heart tissue in the developing embryo.
6.2. Results
6.2.1. CELF1 is expressed in cardiomyocytes and is predominantly nuclear in cardiac
tissue in vivo
CELF1 is highly expressed during vertebrate embryonic development (Ladd
et al., 2001) and its expression in the developing chicken heart is dynamic, as
measured by western blot analysis of whole heart protein extracts (Chapter 4;
Blech-Hermoni et al., 2013). Within the heart, CELF1 is predominantly found in the
133 nuclei of cells in the myocardial cell layer (Chapter 4; Blech-Hermoni et al., 2013;
Brimacombe and Ladd, 2007), as red blood cells and cells in the cardiac cushions and epicardium appear not to express the protein (Chapter 4; Blech-Hermoni et al.,
2013). In order to confirm the identity of the cells expressing CELF1 in the heart, cardiac markers were imaged, in conjunction with CELF1, by immunofluorescence.
Antibodies against Tropomyosin (TPM2), Alpha Actinin 2 (ACTN2), and CELF1 were used in order to visual their expression in chicken whole embryo sections, by indirect immunofluorescence. TPM2 was seen in all cell layers of the heart, while
ACTN2 was found only in the muscle cells (cardiomyocytes), but absent from the endocardial cells lining the luminal surface of the heart or the epicardial cells lining the outer surface of the heart (Figure 17A). CELF1 protein expression was seen in the same cell population as ACTN2, and was not found in endocardial or epicardial cells (Figure 17A). As we have shown before (Chapter 4; Blech-Hermoni et al.,
2013), CELF1 was found predominantly in the nuclei of myocardial cells, in contrast to the cytoplasmic expression of ACTN2 and TPM2 (Figure 17A). These observations confirm that CELF1-expressing cells in the heart are cardiomyocytes and that ACTN2 can be used to identify these cells. In order to evaluate the roles of
CELF1 in embryonic cardiomyocytes, we next optimized a chicken primary embryonic cardiomyocyte culture system for manipulation of CELF1 levels.
13 4
Figure 17. Isolated chicken primary embryonic cardiomyocytes represent a highly enriched cardiomyocyte culture system. Cardiomyocytes in the embryonic day 8 chicken heart in vivo (A) and in culture chicken day 8 embryonic cardiomyocytes (B) express CELF1 protein at high levels in the nucleus as determined by immunofluorescence. Cells of the epicardial (epi) and endocardial (endo) layers of the heart do not express appreciable levels of CELF1 in vivo, while the cultured cells all express CELF1. As in the in vivo tissue, cardiomyocytes continue to express sarcomeric proteins (e.g., ACTN2) and all three cell types express Tropomyosin (TPM2).
135 6.2.2. Cultured chicken primary embryonic cardiomyocytes recapitulate expression of cardiac markers and CELF1 seen in the developing heart
Chicken primary embryonic cardiomyocytes have previously been used to evaluate cardiomyocyte development, physiology, and myofibrillogenesis (DeHaan,
1970; Lin et al., 1989). We have previously used these cells to study the subcellular localization of RBPs, such as CELF2 (Ladd and Cooper, 2004), MBNL1 and MBNL2
(Terenzi and Ladd, 2010). These cells can be isolated to high purities and in great numbers, and are amenable to transfection with plasmids (at low efficiencies) and siRNAs (at high efficiencies). In order to optimize this primary cell culture model for the study of the roles of CELF1 in cardiomyocytes, we evaluated the growth of cells isolated at different embryonic stages and grown under different conditions, and measured expression of CELF1 and cardiac markers.
First, growth of cardiomyocytes isolated from day 8 hearts was compared to that of cardiomyocytes isolated from day 14 hearts, as day 8 (stage 35) was within the developmental window of greatest CELF1 protein expression (Chapter 4), while still yielding sufficient numbers of cells. These cells were grown on Primaria™- treated culture plates (Fisher Scientific), a proprietary treatment that specially modifies the polystyrene surface with to incorporate cationic and anionic functional groups. These plates are recommended for the culture of difficult-to-grow cells, such as a variety of primary cell types, including primary cardiomyocytes. When plated at similar densities, cells isolated from day 8 hearts became confluent more quickly than cells isolated from day 14 hearts, with no other differences being observed. We
136 next evaluated the growth of cells isolated at both developmental stages on different
substrates.
While cell-cell communications are important for the development of the
heart, the extracellular matrix is also essential for proper formation of the tissue and
cardiac function (Bowers et al., 2010; Fomovsky et al., 2010; Lockhart et al., 2011;
Rozario and DeSimone, 2010). These components include both proteins (e.g.,
Collagens, Fibronectin, Laminins) and glycosaminoglycans (Hyaluronan). We tested
growth of cultured primary embryonic cardiomyocytes on different substrates: a mixture of Poly-D-Lysine and Collagen Type I (PDL/COLI), Fibronectin, or
Primaria™-treated culture plates (Fisher Scientific). Plates were coated with
PDL/COLI as follows: PDL (20 µg/mL in water; BD Biosciences) was added to plates
and incubated at room temperature for at least 1 hr, then aspirated; rat tail COLI
(250µg/mL in 0.02N Acetic Acid; BD Biosciences) was then added to the plates,
incubated at room temperature for at least 1 hr, then aspirated; cells were then
plated. Plates were coated with Fibronectin as described in Chapter 2. Following
plating, cells were observed daily. Cells grown on PDL/COLI formed web-like ridges
throughout the plate, an appearance also seen when cells have difficulty adhering to
the plate surface, whereas cells grown on Fibronectin or Primaria™ grew to
confluence and appeared healthy (not shown). Cells isolated from day 8 hearts grew
more vigorously than cells isolated from day 14 hearts, reaching confluence a day
earlier, but otherwise both sets of cells behaved the same (not shown). A significant
difference was observed when CELF1 protein levels were evaluated in the different cell populations over days 5 days in culture, by western blot analysis. After 24 hrs in
137 culture, CELF1 protein levels in both cell populations were low. However, CELF1
protein levels in cells isolated from day 14 hearts continued to decline until, by 72
hours post-plating (hpp), little to no CELF1 protein was detected in these cells (not
shown). By contrast, cells isolated from day 8 hearts expressed higher amounts of
CELF1 protein in the first 48 hrs in culture. However, by 120 hpp, almost no CELF1
protein was detected in cells plated on PDL/COLI or Primaria™, while cells plated on
Fibronectin continued to express CELF1 (Figure 18) and, in fact, CELF1 protein
levels recovered to pre-plating levels (not shown). Following these observations,
only cells isolated from day 8 hearts were used, and cells were cultured on
Fibronectin-coated plates (and on Fibronectin-coated coverslips, as described in
Chapter 2).
Cultured primary cardiomyocytes beat in culture, with spontaneously
beating pacemaker cells (defined as cells exhibiting spontaneous beating in the
absence of contacts with other cells; DeHaan, 1969) being observed by 24 hpp.
Interestingly, cells that were frozen upon isolation and thawed at a later date
resumed beating once plated, and these beating cells could not be visually
distinguished from freshly isolated cardiomyocytes (although expression of
cardiomyocyte markers was not evaluated in these cells). In addition, while
exposure to trypsin (0.13%; see Chapter 2) during the isolation procedure does not
appear to inhibit the cardiomyocytes’ capacity to beat, we found that following
trypsinization (with 0.25% Trypsin) and re-plating, beating was completely lost. We also found that cells ceased to express CELF1 protein following re-plating, with no
138
Figure 18. Expression of CELF1 over days in culture and development of myofibrillar structure. (A) CELF1 levels as measured by western blot analysis are low shortly after plating (24 hpp), but pre-plating levels of expression (not shown) are recovered quickly (48 hpp). (B) Reassembly of the myofibril is complete within 48 to 96 hours of plating as shown by immunofluorescence using sarcomeric markers (e.g., ACTN2 and TTN). Note that the shape and size of the cells changes over days in culture, in addition to the myofibrillar organization: upon plating, cells are small and rounded, and they become flatter and larger over days in culture (the TTN-stained cell at 96 hpp illustrates the bi-nucleate state of many of the cardiomyocytes as they are cultured for prolonged periods of time). Fluorescent images were taken at the same magnification.
139 qualitative effect on viability or proliferation (data not shown). Only freshly isolated cells were cultured for all experiments described here.
Finally, the identity of the cultured chicken primary embryonic cardiomyocytes was further validated by comparing expression of cardiac markers and of CELF1 in the cultured cells to their expression in the embryonic heart. Using the same antibodies as described above, cultured cells were probed for ACTN2,
TPM2, and CELF1. As expected, all cells were found to express TPM2 (Figure 17B), consistent with its ubiquitous expression in the chicken embryonic heart (Figure
17A). The vast majority of the cultured cells showed ACTN2 expression, indicating that the culture was very pure, and the expression pattern revealed striated fibers consistent with myofibrillar staining (Figures 17B and 18). CELF1 was found to be
expressed in all the cultured cells, further supporting the observation that this
culture system represents a highly purified myocardial cell population. Finally,
CELF1 expression was observed to be predominantly nuclear by immunofluorescence (confirmed by nuclear counterstaining with DAPI), in line with our observations of the expression of CELF1 in the myocardium of the embryonic heart (Figure 17A; Blech-Hermoni et al., 2013). Taken together, these findings indicate that these cells provide a cell culture system that correctly recapitulates
expression patterns of CELF1 and of cardiac markers seen in the embryonic heart at
stages relevant for the study of cardiac morphogenesis.
140 6.2.3. Knockdown of CELF1 in cultured primary embryonic cardiomyocytes leads to myofibrillar disorganization
In order to investigate the role of CELF1 in embryonic cardiomyocytes,
CELF1 protein levels were knocked down by siRNA-mediated degradation of transcripts. Transfection of cultured cells with two independent siRNAs (si1 and si2) designed to target the chicken CELF1 transcript resulted in robust knockdown of CELF1 in cultured embryonic primary cardiomyocytes, by western blot analysis
(as previously shown; Figure 15) and by immunofluorescence (Figure 19). A control (non-targeting) siRNA (siCont) did not result in changes to CELF1 protein or transcript levels. Following knockdown of CELF1, the organization of the contractile apparatus was evaluated by immunofluorescence. Antibodies against components of the Z-discs (ACTN2 and TTN) and the M-lines (MYOM)(Figure 20B) were used. All sarcomeric markers tested revealed disorganization following knockdown of CELF1: while mock- and siCont-treated cells contained longitudinal, thick myofibrils, as indicated by long striations, si1- and si2-treated cells contained thin, often web-like myofibrils with small, globular striations (Figure 20B). While cell size was not evaluated in these cells, the cells did not appear markedly different under brightfield microscopy (not shown). Importantly, transfected cardiomyocytes continue to beat in culture, and beating in si1- or si2-transfected cells was indistinguishable from beating in mock- or siCont-transfected cells by visual inspection of video microscopy (not shown).
141
Figure 19. Transfection of cultured chicken primary embryonic cardiomyocytes with siRNAs results in robust knockdown of CELF1 protein and transcript levels. Depletion of CELF1 with two independent siRNAs (si1 and si2) but not a control siRNA (siCont) can be seen by western blot (A; GAPDH, loading control) and by immunofluorescence (B). Knockdown of CELF1 leads to reductions in both nuclear and cytoplasmic CELF1. (C) Transfection of cells with siRNAs results in robust knockdown of CELF1 transcript levels, by real-time RT-PCR (values were normalized to GUSB levels; *, p < 0.05; **, p < 0.01; t-test versus M). Cells were transfected at 24 hpp and collected at 72 hpt.
142
Figure 20. Knockdown of CELF1 using siRNAs leads to reduced cell proliferation and profound disorganization of myofibrillar structure in cultured chicken primary embryonic cardiomyocytes. (A) Knockdown of CELF1 (with si2) at 24 hpp leads to a reduction in proliferation 48 hpt, as measured by EdU incorporation (*, p < 0.05; one-tailed t-test against siCont). (B) Knockdown of CELF1 at 24 hpp leads to disruption of myofibril organization (by visualizing ACTN2, TTN, and MYOM immunofluorescence) at 72 hpt.
143 As CELF1 has been implicated in cell-cycle arrest and apoptosis (Chang et al.,
2012; Liu et al., 2014; Talwar et al., 2013; Xiao et al., 2011), proliferation of these cultured cardiomyocytes was also evaluated by incorporation of a nucleoside analog, 5-ethynyl-2’-deoxyuridine (EdU), during replication of the DNA in the S stage of the cell cycle. Under the growth regimens employed in the experiments described thus far, cardiomyocytes became confluent within 48-72 hpp (24-48 hpt) and, once a monolayer was formed, proliferation slowed down. In order to evaluate proliferation, cells needed to be of low enough densities that contact inhibition did not present a concern. Since cells were routinely transfected at 24 hpp, we chose to plate cells at lower densities and evaluate them at an earlier time point, thereby balancing the proliferation rate with the need for effective knockdown. Cells were thus plated onto 60 mm culture plates (resulting in one third the usual density of cells per plate), transfected at 24 hpp, and evaluated at 48 hpt. Robust knockdown was achieved under these conditions (not shown). Treatment of cells with siRNAs against CELF1 resulted in a dose-dependent reduction in proliferation, with a robust effect seen following treatment with the siRNA that consistently produced greater levels of CELF1 knockdown (si2; Figure 20A) and a trend toward reduction with
the less effective siRNA (si1), that did not attain statistical significance.
6.2.4. CELF1 is required for early events in myofibril assembly in cultured cells
During isolation of the cardiomyocytes, myofibrils disassemble and must be reassembled once the cells begin making contacts with the plate surface and with
144 other cells (Lin et al., 1989). The precise reasons for this are not fully understood,
but physical forces to which the cells are subjected to during isolation, as well as
chemical agents (such as trypsin; Legato, 1972), have been suggested to play a role.
Since assembled myofibrils are necessary for beating to occur, the fact that beating is observed by 24 hpp suggests that this process can be rapid. However, the ability of cardiomyocytes to contract also depends on their permeability to ions that promote contraction (such as Ca2+), as well as their sensitivity to inhibitory ions
(such as K+). In a population of cultured embryonic cardiomyocytes, some cells are
more sensitive to extracellular K+ while others are less so, and this proportion
changes during development (DeHaan, 1970). The fact that only some of the cells
could be seen to beat at this early stage is in line with observation that only a subset
of isolated cells can contract spontaneously in the absence of contacts with
neighboring cells (pacemaker cells; DeHaan, 1968). By 2-3 days in culture, cells had
proliferated to near confluence and beating could be seen in large synchronized foci
throughout the culture dish.
The myofibril assembles in a stepwise process (Gregorio and Antin, 2000). In
order to determine whether the knockdown of CELF1 (seen as early as 24 hpt;
Figure 21A) prevents or delays the assembly of the myofibril, or if it leads to
disassembly of assembled myofibrils, we imaged the myofibrils at consecutive days
in culture, from 24 hpt to 96 hpt, and then compared the appearance of the normally
assembling myofibrils to that of the myofibrils following CELF1 kncockdown
145
146 Figure 21. Following knockdown of CELF1 at 24 hpp, myofibrillar disorganization persists. (A) Knockdown of CELF1 protein persists for as long as 96 hours post-transfection, by western blot analysis (Ponceau S staining shows protein integrity and relative levels of total protein). (B) Myofibrillar organization was monitored by immunofluorescence using sarcomeric markers (ACTN2 and MYOM) over 96 hours following mock (M) or siRNA (si2) transfection. Cells were transfected at 24 hpp and fixed at 24, 48, 72, and 96 hpt.
147 (Figure 21B). By evaluating the distribution of myofibrillar protein, we found that different components appear to become organized at different time points: myofibrils visualized using markers of the Z-discs (ACTN2 and TTN) became progressively thicker and more organized until 72 hpt, after which time no further change was observed; during the same time course, myofibrils visualized using a marker of the M-lines (MYOM) appeared well organized by 24-48 hpt and no further change in the appearance of the myofibrils was observed (mock-treated [M] panels in Figure 21B). These observations are in agreement with earlier studies of the progression of myofibril reassembly in cultured primary avian cardiomyocytes
(Komiyama et al., 1990; Terai et al., 1989). In addition to the state of the myofibril, it is important to keep in mind that the cells also undergo changes in size and shape during this time course: at 24 hpt, cells were small and either ovoid or spindle- shaped, while after that time they became flatter and wider. The combination of these two factors resulted in staining that appeared punctate in the cells at 24 hpt
(with the exception of MYOM, which already exhibited striated expression even at this early time point), appeared as globular beads on thin myofibrillar strings at 48 hpt, and then exhibited a mature-looking appearance at 72 hpt. Following knockdown of CELF1, the myofibrils at 24 hpt appear as unassembled as the myofibrils in the mock-treated cells. However, while the myofibrils in the mock- treated cells become progressively more assembled, the myofibrils in the si2- treated cells become longer, and sometimes assume a web-like appearance (si2- treated panels in Figure 21B). At no time point did the striation become more rod- shaped, which would suggest maturation of the myofibril. This may indicate that,
148 while the cells continue to grow and divide following knockdown, the myofibrils
retain an immature organizational state.
Next, we attempted to evaluate if recovery of myofibrillar organization would
occur once the siRNA-mediated knockdown of CELF1 had dissipated, presumably leading to recovery of CELF1 protein levels to normal. It should be noted that although the growth medium was changed every two days in culture (allowing for
the siRNAs to be washed out of the culture), CELF1 protein levels were not observed
to recover to normal levels even at 96 hpt (Figure 21A), and up to 144 hpt (not
shown). ACTN2 and TTN failed to fully attain an assembled appearance even by 96 hpt (Figure 21B). During the same time course, myofibrils stained for MYOM appear no more assembled at 96 hpt than they are at 24 hpt, with myofibrils in many cells appearing less organized following CELF1 knockdown (Figure 21B).
These observations support a role for CELF1 in the assembly or organization of the myofibril. One possibility that would explain the difference between the organization of Z-disc markers (ACTN2, TTN) versus an M-line marker (MYOM) is that CELF1 may regulate one key factor in the organization (or assembly) of the myofibril, but different components are affected differently due to their different kinetics of incorporation. Alternatively, CELF1 may regulate several different nodes in myofibril assembly or maintenance, and different markers each illuminate a small part of the picture. These possibilities should be tested in future investigations.
Finally, the effect of CELF1 knockdown on the level of myofibrillar organization appeared to depend on the efficiency of CELF1 depletion: cells in which
CELF1 was depleted using si1 exhibited myofibrillar disorganization less severe
149
150 Figure 22. Developmental expression and activity of Celf1 are conserved in Xenopus laevis. Levels of Celf1 protein were evaluated in whole embryo extracts (A; stage 24 through stage 49) and in isolated hearts (B; stage 32 through adult) by western blot analysis; Ponceau S staining was used to indicate equal loading. (C) celf1 transcript levels were evaluated in embryonic (stage 32 through stage 49) and adult hearts (n=3-4) by real-time RT-PCR. Real- time RT-PCR primers (Table 4) were designed to amplify both celf1a and celf1b. Error bars indicate standard error of the mean; (D) Intracellular localization of Celf1 was visualized by immunofluorescence in whole heart (top; Actin, phalloidin staining; 3D reconstruction of a confocal stack) and in cryosections (bottom; scale bar = 100μm). (E) The ability of frog Celf1 to regulate alternative splicing was assayed in COSM6 (Green African Monkey kidney epithelium) cells. The CELF-responsive RTB33.51 mini-gene was transfected into COSM6 cells along with celf1a at 24 hpp, RNA was collected at 72 hpt, and spliced mini-gene transcripts were amplified by radiolabeled RT-PCR and quantitated (top). Exogenous protein expression was confirmed by western blot analysis (WB). Lanes in panel E come from the same gels, while intervening lanes were removed for clarity. One-tailed t-tests were applied in panels C (against stage 32) and E (against RTB33.51-only); **, p < 0.01.
151 than that seen in si2-treated cells (Figure 20B). Taking together these findings and
the dose-dependent effect of CELF1 knockdown on cardiomyocyte proliferation
suggests that CELF1 is regulating these systems along a gradient of activity.
6.2.5. CELF1 expression patterns are conserved between frog and chicken
In order to further investigate the effect of CELF1 knockdown on the
cardiomyocytes of the developing heart in vivo, we adopted the African Clawed Frog
(Xenopus laevis) as a model organism. The frog is a common model organism for
studies of embryonic development, although its use in the study of heart
development has been largely relegated to early events of tissue specification,
patterning, and inductive signals. Nevertheless, X. laevis provides various advantages for the study of cardiac morphogenesis, including the fact that morpholino microinjection can be easily used to induce protein knockdown and that within the time course of interest, diffusion of oxygen from the environment and the presence of intracellular nutrient pools allow the embryo to develop normally in the absence of a beating heart (Blitz et al., 2006). Levels of Celf1 transcripts have been
evaluated in whole embryo (Wu et al., 2010b), but protein levels have never been
measured, and neither transcript nor protein has been measured in the developing
heart. We therefore compared protein levels in whole embryos and in isolated
hearts from a series of stages. We found that Celf1 levels in whole embryo gradually
increased during development (Figure 22A), which is consistent with Celf1
transcript levels measured in whole embryo, at least between stage 11 and stage 43
152 (Wu et al., 2010b). However, Celf1 levels in isolated hearts underwent a dramatic
increase during embryogenesis, peaking at stage 46 before steeply declining at stage
49 and remaining low in adult heart tissue (Figure 22B). Unlike in whole embryo,
transcript levels did not change in the same way, but rather remained largely
unchanged (Figure 22C), with one exception: between stage 46 and stage 49, transcript levels decline precipitously (and statistically significantly, compared with stage 32), and this remain significantly reduced in adult heart. This suggests that
Celf1 may be regulated post-transcriptionally during stages of cardiac morphogenesis (stage 32 to stage 46), as we have previously shown in the chicken
(Chapter 4; Blech-Hermoni et al., 2013), while after this stage changes in transcript and protein levels become more concordant, suggesting that contribution of transcriptional regulation cannot be excluded. Currently, only one report has been published showing transcriptional regulation of CELF1 expression, showing transcriptional activation by CBP/p300/E12 and myogenin/p300/E12 complexes in myoblasts and myotubes, respectively (Huichalaf et al., 2007), but little is known about regulation of CELF1 expression in the heart.
Tissue distribution of celf1 transcripts has also been evaluated in the developing frog embryo (Wu et al., 2010b), but the heart was not analyzed. Since we found that Celf1 protein and transcript levels in the heart were not concordant during cardiac morphogenesis, we chose to visualize Celf1 protein using indirect immunofluorescence. As we have seen in the chicken and mouse embryos (Chapter
4; Blech-Hermoni et al., 2013), Celf1 was detected in muscle tissue (both cardiac and skeletal muscle, including the somites and the hypaxial muscle clusters), the eye,
153 and the neural tube (not shown). Expression of Celf1 in muscle cells was
predominantly nuclear (Figure 22D and data not shown), while expression in eye and neural tube was predominantly cytoplasmic (not shown). In the heart, Celf1 expression was found to be restricted to the nuclei of cardiomyocytes, as demonstrated by immunofluorescence in embryonic whole heart (stage 47; three- dimensional reconstruction of a confocal stack) and in sections through heart wall
(stage 41; epifluorescence)(Figure 22D).
Finally, we asked whether frog Celf1 could regulate alternative splicing. The open reading frame of the frog celf1a was amplified from stage 41 frog heart total
RNA and cloned into the pCS2+ expression vector. A mini-gene reporter was used to evaluate the ability of Celf1a to promote inclusion of a CELF-responsive alternative exon. The RTB33.51 mini-gene (described by Ryan and Cooper, 1996) directs the transcription of a pre-mRNA comprised of an alternative exon (TNNT2 exon 5;
described in Chapter 1), flanked by constitutive exons and the intervening introns.
Cis regulatory elements within the proximal introns promote CELF1-mediated
enhancement of exon inclusion. The expressed RNA is not translated and inclusion
of the alternative exon can be measured by RT-PCR with primers in the flanking
exons. This mini-gene exon was shown to be sensitive to regulation by human
CELF1 (Ladd et al., 2001) and chicken CELF1 (Ladd et al., 2005a). COSM6 cells
normally express very low levels of endogenous CELF1, and so these cells were
chosen to evaluate the activity of exogenously expressed protein. Cells were
transfected with the frog Celf1a expression construct (pCS2-Celf1a; described in
Chapter 2) along with the RTB33.51 reporter (pCS2-RTB33.51; described in
154 Chapter 2), RNA was collected 48 hrs post-transfection, and RT-PCR was used to evaluate exon inclusion. While COSM6 cells transfected only with the mini-gene construct exhibited a lack of CELF1-mediated activity, as demonstrated by the predominance of the exon-skipped RTB33.51 variant, co-transfection of cells with the pCS2-Celf1a construct resulted in a switch in splicing from nearly 70% skipping to roughly 70% inclusion of the alternative exon (Figure 22E).
Taken together, the expression patterns and splicing activity of frog CELF1 recapitulate our finding in the chicken and mouse hearts during embryonic development (Chapter 4; Blech-Hermoni et al., 2013), demonstrating conservation of CELF1 regulation between mammals, birds, and amphibians during
cardiogenesis.
6.2.6. Knockdown of Celf1 in frog embryos results in cardiac looping defects and
cardiac dysmorphia
Establishing asymmetry is an active, evolutionarily conserved, process
(‘symmetry breaking’) requiring various chemical and structural components
(Palmer, 2004). The heart precedes all other organs in developing morphological asymmetry, and this ‘handedness’ is essential for both its form and its function
(Manner, 2013). In the straight heart tube, peristaltic contraction is sufficient for transporting the relatively small volume of blood around the early embryo. As the embryo grows, the blood volume increases and so must the size of the heart and its efficiency. In order to create an efficient pump within the spatial confines of the
155
156 Figure 23. Morpholino-mediated knockdown of celf1 in Xenopus laevis leads to cardiac looping defects, ventral edema, and gut malformation. 2-4-cell embryos were injected with Celf1-specific morpholino oligonucleotides and Celf1 protein levels were then evaluated by western blot analysis (A) at stage 35-36 (STD-MO, control morpholino, Celf1-MOs, a mix of two Celf1-targeting morpholinos). (B) Cardiac looping was evaluated in injected embryos (stage 35-36) by whole-mount immunohistochemistry using antibodies against meromyosin (MF20) or TNNT2 (CT3). Hearts are outlines in broken white lines; a path through the middle of the heart tube is indicated with a solid black line. D-loop, dextral-loop; L-loop, levo-loop. (C) Embryos were collected at stage 46 and evaluated for morphological defects by light microscopy of whole embryos (filled arrowhead indicates malformed gut; open arrowhead indicates ventral edema). (D) Embryos were collected at stage 46 and evaluated for morphological defects by histological staining (Hematoxylin & Eosin) of paraffin embedded sections (V, ventricle; OFT, outflow tract); 10 µm section thickness; scale bar = 100 µm; n=2.
157 pericardial cavity, the chambers are expanded, internal ridges and constrictions are
formed to more precisely and forcefully direct the blood, and angles are introduced
to the tube (Moorman and Christoffels, 2003). This is achieved by a highly coordinated and reproducible right-handed kinking (or dextral looping; D-looping)
of the heart tube, through a C-shaped intermediate to an S-shaped structure
(Manner, 2000). In tetrapods, this process also positions the ventricles and atria
such that the appropriate chambers open into each other, and such that the large
vessels are correctly placed and oriented.
In order to evaluate the effects of Celf1 knockdown on the developing heart, we
microinjected 2- to 4-cell embryos with two morpholino oligonucleotides (MOs)
designed to target the translation start site of both frog CELF1 co-orthologs (Celf1a
and Celf1b; EDEN-BP MOs described by Gautier-Courteille et al., 2004). We chose to
evaluate cardiac looping for two reasons: 1. Looping is a robustly reproducible,
externally visible, hallmark of cardiac morphogenesis, which involves the
remodeling of cardiomyocyte populations, and 2. Disruption of celf1 levels in
zebrafish (Danio rerio) was shown to lead to defects in somite symmetry as well as
cardiac looping (Matsui et al., 2012). Microinjection of MOs (6.4 pmol/embryo)
resulted in robust knockdown of Celf1 at stage 35-36, and also at stage 46, as
measured by western blot analysis (18.7% remaining at stage 35-36 and 58.5%
remaining at stage 46; Figure 23A). A standard control MO (STD-MO) had little
effect on Celf1 levels.
Following microinjection of MOs, embryos were collected at stage 35-36 and
immunostained using antibodies either against a myosin heavy chain (Meromyosin;
158 MF20) or against Tnnt2, in order to highlight the heart. While uninjected embryos predominantly exhibited a normally right-looped heart (D-loop), and embryos microinjected with a non-targeting control MO showed few instances of cardiac abnormalities, in embryos microinjected with Celf1-targeting MOs (Celf1-MOs) both cardiac looping and cardiac morphology were abnormal (Figure 23B). In uninjected and STD-MO-treated embryos, hearts were rarely observed that were unlooped
(2/185 and 4/97, respectively) or incorrectly looped (0/185 and 3/97, respectively). However, these were quite commonly seen in Celf1-MOs-treated embryos (41/139 and 10/139; p < 9E−06 for both phenotypes, versus uninjected;
Fisher exact test). In fact, while about half the hearts observed in Celf1-MOs-injected embryos were looped in the correct direction (D-looped), the degree of looping appeared less pronounced than observed in D-looped hearts in uninjected or STD-
MO-injected embryos. The correctly looped hearts in Celf1-MOs-injected embryos appeared stouter than normal hearts and indeed suggested an intermediate state between correctly looped and unloooped hearts. Furthermore, the incidence of cardia bifida, which is not a looping defect but rather a failure of fusion of the bilateral heart fields (Brand, 2003), was also higher in this group (11/141 in Celf1-
MOs-injected embryos versus 3/186 in uninjected embryos; p = 0.0104; Fisher exact test). In order to evaluate how these looping defects may affect the embryos at a later stage of development, embryos were allowed to grow to stage 46, by which time cardiac morphogenesis is complete in uninjected embryos. Embryos were then collected, fix, and paraffin sections were stained for histological analysis. By evaluating several treated embryos, we found various levels
159 of morphological abnormality in the hearts of Celf1-MOs-injected embryos, including chamber dilation, thinning of the chamber wall and reduced trabeculation
(Figure 23D). These features are reminiscent of the dilation seen in the MHC-CELFΔ mice, in which CELF activity is repressed in the hearts of young mice (Ladd et al.,
2005b). While functional parameters of cardiac contraction and contractility were not directly evaluated in injected embryos, ventral edema was observed in Celf1-
MOs-injected embryos (Figure 23C), which suggests that cardiac dysfunction may be present.
Finally, embryos injected with Celf1-MOs exhibited gut dysmorphia (Figure
23C). This phenotype is less severe than what would be expected, base on findings in zebrafish that show that knockdown of celf1 in the embryo results in loss of some endoderm-derived organs (e.g., liver), reduced endodermal cell proliferation, and reduced endodermal cell migration (Tahara et al., 2013). This group suggested that celf1 regulates endoderm-derived organogenesis by regulating targets that control cell growth (gata5) and migration (cdc42). Our findings are also in line with studies of the establishment (or maintenance) of both bilateral symmetry patterning, which found that both knockdown and over-expression of CELF1 in zebrafish and frog embryos results in somite segmentation asymmetry (a defect of bilateral body symmetry patterning) and randomization of cardiac looping (a defect of proper left- right asymmetry patterning)(Cibois et al., 2013; Cibois et al., 2010; Gautier-
Courteille et al., 2004; Matsui et al., 2012).
160
Figure 24. Morpholino-mediated knockdown of Celf1 in Xenopus laevis leads to myofibril disorganization in the developing heart. Myofibrillar organization was evaluated in heart wall of stage 46 embryos by immunofluorescence (as in cultured cardiomyocytes), following MO microinjection. In uninjected embryos (A), mature myofibrils are visible as interdigitating Actn2 and Actin staining, whereas Actn2 staining is more diffuse in Celf1-MOs-injected embryos (B), and does not associate with actin striations in the myofibrils. Top panels show the separate green and red channels for the indicated box in the merged image (Actin, phalloidin staining). 7 µm section thickness; scale bar = 20 µm; n=2.
161 Finally, the myofibrillar organization in cardiomyocytes of the embryonic
frog heart was evaluated by immunofluorescence. Embryos were injected with
Celf1-MOs, or were left uninjected, and were collected at stage 46. Cryosections
were subjected to indirect immunofluorescence probing for Actn2 and filamentous
Actin (using fluorophore-conjugated Phalloidin). In heart wall from uninjected
embryos, myofibrils are readily visible and striations are prominent, with Actn2
(green) and Actin (red) interdigitating along the length of the myofibrils (Figure
24A; small panels show the separate red and green channels, while the large panel
shows the merged image). By contrast, in Celf1-MOs-injected embryos, myofibrils
were sparse within the cardiomyocytes and Actn2 staining was more diffuse, and punctate in some regions, while it was largely not seen to interdigitate with Actin staining along myofibrils (Figure 24B). These findings support our observation in cultured chicken primary embryonic cardiomyocytes. As in the cultured cells, hearts
of injected embryos continue to beat, even as the contractile apparatus in their
cardiomyocytes appear do be disorganized.
6.3. Discussion
The Pre-myofibril Model of myofibril assembly posits that fiber rudiments
(‘pre-myofibrils’) come together and are assembled into mature myofibrils over
time (reviewed in Sanger et al., 2010), and a similar process is undergone in
cultured pre-cardiac explants (Gregorio and Antin, 2000; Rudy et al., 2001) and
cultured cardiomyocytes following isolation. The repression of cardiac CELF activity
in young mice results in contractile defects and dilated cardiomyopathy (Ladd et al.,
162 2005b; Terenzi et al., 2009), suggesting a role for this protein in the regulation of the
contractile apparatus post-natally. In order to investigate the role of CELF1 in
myocardial cells of the developing heart, we adapted a primary embryonic
cardiomyocyte culture system and evaluated myofibrillar organization following
CELF1 ablation. Following siRNA-mediated knockdown of CELF1 in cultured primary cardiomyocytes, we observed widespread myofibrillar disorganization.
Furthermore, when we compared the early stages of myofibril assembly in the cultured cells to the disorganization following knockdown of CELF1, we found that the appearance of the myofibrils (i.e., thin fibers punctuated with globular-looking striations) were not dissimilar from the assembling myofibrils at early stages.
However, the size and shape of the cells during these early steps of myofibril assembly is dramatically different from those of the treated cells, as all the cultured cells begin as rounded or spindle-shaped and progressively adopt a flattened, spread-out morphology. This may suggest that the assembly of the myofibrils is slowed down following knockdown, or it may indicate that assembly is completely halted. While we attempted to evaluate if the myofibrils would resume assembly once the effect of the siRNA-mediated knockdown wore off, we were not able to observe recovery of CELF1 protein levels (as long as five days after transfection).
These findings suggested that CELF1 involvement in myofibrillar organization is intrinsic to cardiomyocytes. However, in order to evaluate if myofibril disorganization could also be detected in the developing tissue in vivo, we knocked
down Celf1 in the developing frog embryo. The same phenotype was observed in
Celf1-MO-injected embryo as we had observed in the cultured primary
163 cardiomyocytes, suggesting that Celf1 is necessary for the proper assembly of the
myofibril in the cardiomyocytes in vivo.
In skeletal myoblasts, CELF1 has been implicated in both proliferation and
differentiation into myotubes (Salisbury et al., 2008). CELF1 has been shown to
regulate the translation of p21, a cyclin-dependent kinase inhibitor that restricts re- entry into the cell cycle, and increased nuclear relocalized of CELF1 in DM patient muscle cells leads to enhanced proliferation of myocytes and negatively affects their differentiation state (Iakova et al., 2004; Timchenko et al., 2001b). Our results are in agreement with these findings, as cultured embryonic cardiomyocytes are normally proliferative and express high levels of CELF1. Following knockdown of CELF1, we observe dose dependent reductions in proliferation, suggesting that the cells have exited the cell cycle. Knockdown of celf1 was also shown to lead to reduced proliferation in endodermal cells during zebrafish embryonic development (Tahara et al., 2013), suggesting a general involvement in this biological process. In fact, competition between CELF1 and Calreticulin, CRT, has been suggested to make up a biological switch between senescence and proliferation in hepatocytes and fibroblasts (Barreau et al., 2006; Iakova et al., 2004; Timchenko et al., 2005).
In zebrafish, both knockdown and over-expression of celf1 result in left-right patterning defects, resulting in somite asymmetry and cardiac looping abnormalities
(Matsui et al., 2012). Binding of celf1 to the 3’ UTR of dmrt2a transcripts leads to their degradation, and reductions in transcription factor dmrt2a help to explain the observed patterning defects. However, overexpression of celf1 in embryos also leads to symmetry defects, suggesting that celf1 may have additional targets
164 involved in this cellular process. Levels higher than those shown to be sufficient to
produce symmetry defects result in further abnormalities in different
developmental processes, leading the authors of the study to suggest that celf1 may
have different thresholds for different targets (Matsui et al., 2012). We find that knockdown of celf1 results in cardiac looping defects and other morphological defects in the heart, suggesting that the involvement of celf1 in this process is evolutionarily conserved.
The same group also reported defects in symmetry patterning of endoderm- derived organs, such as liver and pancreas, following knockdown of celf1 (Tahara et al., 2013). Partly, these findings are confounded by the fact that knocking down celf1 resulted in reductions in both proliferation of endodermal cells and in their migration speeds, which may give rise to under-developed or diminutive organs.
However, they also observe a failure of gut looping, which is unlikely to be due to its cellularization alone. In our study in the developing frog embryo, we also observe gut morphological abnormalities following knockdown of Celf1. Since the heart is mesoderm-derived, these findings suggest that Celf1 regulates factors that more generally control the establishment of symmetry in the embryo.
6.4. Acknowledgements
We would like to thank Dr. Debora Malta Cerquiera Santos, Uyen Tran, and
Jessie M. Hassey for their help with maintaining the frogs and for technical assistance. We would also like to thank Dr. Judith Drazba and Dr. John Peterson for
their technical support and advice in confocal imaging. This work was supported by
165 a grant to A.N.L. from the National Institutes of Health (1R01HL089376) and a grant to O.W. from the National Institutes of Health (5R01DK080745-05). Y.B-H. was supported in part by a National Institutes of Health training grant
(5T32GM008056).
166 Chapter 7: Concluding remarks
7.1. Expression patterns and activities of CELF proteins
In Chapter 4, the expression of CELF1 and CELF2 in different tissues of the
developing chicken and mouse embryos is described. The presence of tissue-
specific, as well as cellular-compartment-specific, isoforms of these proteins is
discussed. Little is currently known about the function or consequences of having
this broad array of isoforms, or even their identities. Phosphorylation of CELF1
affects its localization, stability, and activity (Roberts et al., 1997; Timchenko et al.,
1996a). To date, CELF1 has been shown to be phosphorylated by DMPK (Roberts et
al., 1997), PKC (Kuyumcu-Martinez et al., 2007), Akt (Salisbury et al., 2008), and
Cdk4 (Salisbury et al., 2008), but the array of isoforms we observe for two of the six
CELF members suggest that these modifications may not be the only contributors. In fact, both post-transcriptional and post-translational modifications may interact to underlie this diversity. CELF1 and CELF2 are known to undergo alternative splicing
(Choi et al., 1999; Li et al., 2001; Suzuki et al., 2012; Takahashi et al., 2000;
Takahashi et al., 2001) and a variety of CELF2 variants are found that are generated through alternative promoter usage (Ramalingam et al., 2008). In our own studies, we identified several splice variants of both CELF1 and CELF2 transcripts (not shown), although the presence of these variants did not appear to correlate with specific bands observed by western blot analysis (Chapter 4). Further studies designed to uncover the identities of the different variants of the CELF members are necessary in order to elucidate the tissue- and developmental stage-specific
167 functions of these proteins. Seeing as the members of this protein family show a great breadth of expression patterns, distributions, and functions, further
investigation is essential in order to understand the myriad roles of these proteins.
It has been shown that CELF1 and MBNL1 (another RBP that is also found in
the developing heart) can regulate each other, presumably through mutual binding sites in their 3’ UTRs (Masuda et al., 2012), and these proteins have been proposed to regulate (often antagonistically) some of the same targets. It is quite likely that there is overlap in targets of the different CELF proteins, but especially CELF1 and
CELF2 (which are found in many of the same tissues). It is not currently known in what way these proteins are maintained functionally distinct. CELF1 and CELF2 are both found in the developing heart and future studies should aim to elucidate if they have completely unique sets of targets in vivo, or if there is indeed overlap. In
Chapter 5, we demonstrate that MYH7B is regulated by CELF1, both directly and
indirectly. It is intriguing to speculate that this kind of complex regulation is also
employed by different members of the family to regulate the expression of shared
targets. Unlike the example of MYH7B, of course, different members can regulate
shared targets in opposite ways, directly or through intermediaries. In fact, the
study of regulation by RBPs often aims to identify direct targets. Our finding that
Myh7b can also be regulated indirectly through splicing-mediated reduction of
Mef2a transcriptional activity suggests that the magnitude of regulation by these
proteins can extend well beyond the individual targets. Our analysis of direct
binding sites also indicates that binding often occurs in the middle of very large
introns, far from annotated splice sites. While some proximal splice sites may simply
168 not be annotated, this observation does raise the possibility that either CELF1 has
yet unknown functions that are mediated by binding at these distant sites, or that
binding at these sites represents capture of low affinity interactions that occur in
many places along the transcriptome (seeing as GRE-like sequences are not rare,
even by chance). Another possibility is that CELF proteins regulate common targets
in common ways, and so there is a need to maintain a certain ‘appropriate’ level of
CELF activity for the cells to function appropriately.
7.2. CELF1 in myofibrillar organization and cardiac morphogenesis
In Chapter 6, we investigate the phenotypic effect of CELF1 knockdown in
embryonic cardiomyocytes in culture and in the developing heart. We observe that
the organization of various components of the sarcomere into myofibrils is
disrupted and we present data that suggest that this disorganization represents a
developmental delay or arrest, rather than a structural disassembly. In the process of muscle differentiation, the production of muscle-specific proteins must be initiated before structures such as the myofibril can be assembled, and CELF1 could
play a role in this sequence of events in the developing heart. For example, Cardiac
Troponin T, which is a component of the Troponin Complex that transduces
chemical signals (i.e., cytoplasmic Ca2+ release) into physical force (i.e., contraction),
is alternatively spliced to give rise to several isoforms in human heart (Adamcova
and Pelouch, 1999). Expression of these isoforms in the myofibril peaks at different
times in development, endowing the myofibrils with different properties. For
instance, isoforms 1 and 2 are expressed in the fetal heart, while isoform 3 is
169 predominantly expressed in the adult heart. TNNT2 isoforms 1 and 2 contain
residues that increase their sensitivity to Ca2+, which is suggested to be important in the fetal heart, where amplitudes of Ca2+ fluxes are low (Adamcova and Pelouch,
1999). Thus, regulating alternative splicing of components of the myofibril could have dramatic effects on function. In a similar way, it is intriguing to speculate that altering the interacting domains of a myofibrillar structural protein may affect its ability to incorporate into the assembling fiber. Components of the myofibril can recycle rapidly in and out of the myofibril (Sanger et al., 2010 J Biomed Biotech), providing a powerful platform for regulation of contractile function. The cultured cardiomyocytes in our studies were isolated from day 8 hearts, by which time these cells already contain functional myofibrils. Once these cells are plated, we do notice that CELF1 levels are briefly reduced (Figure 18), before recovering to normal levels, coincident with the reassembly of the myofibrils (data not shown). This suggests that the isolation procedure may impact the levels of normally expressed proteins, and that knockdown of CELF1 may somehow impede the recovery of these proteins to normal levels, or interfere with their assembly. CELF1 has been extensively studied in its capacity as a regulator of alternative splicing, and especially how its abnormal activation in DM1 results in a reversal of the “fetal to adult” transition in the splicing programs of various target genes. In the MHC-CELFΔ mouse (Ladd et al., 2005b), which transgenically expresses a dominant negative
CELF protein (Charlet-B. et al., 2002a) in the myocardium, CELF nuclear activity is specifically repressed in the hearts of the mice starting at about 2 weeks post- partum. These mice exhibit profound dilated cardiomyopathy, cardiac hypertrophy,
170 contractile defects, loss of myofibers, and fibrosis. Furthermore, the loss of CELF
protein nuclear activity in these mice results in mis-splicing of several known
targets of CELF1 alternative splicing (e.g., Mtmr1, Mef2a, Itgb1, and Bin1), of which
two are known to be involved in aspects of myogenesis (MEF2A) and contraction
(ITGB1). However, by 2 weeks post-partum, CELF1 levels are normally very low in
the heart, underscoring the essential role of CELF proteins in heart tissue.
Importantly, different targets of CELF activity are important at different stages of
cardiac development. Of the two MHC-CELFΔ mouse lines reported, one expresses
higher levels of transgenic dominant negative protein (MHC-CELFΔ-10) in the heart and exhibits more severe pathology, while the other expresses lower levels of dominant negative protein (MHC-CELFΔ-574) and exhibits a more mild phenotype
(Ladd et al., 2005b). Mice of the MHC-CELFΔ-574 line exhibit cardiac pathology,
which resolves over time, and abnormal splicing, which does not (Terenzi et al.,
2009).
In Chapter 5, we discuss the identification of MYH7B as a direct target of
CELF1 through CLIP analysis in hearts from day 8 chicken embryos. We describe
how MYH7B is regulated by CELF1 both directly (through the splicing of a
previously unannotated exon, which likely leads to degradation of the transcript)
and indirectly (through CELF1-mediated splicing of Mef2a, which reduced its
transcriptional activation of Myh7b). How the altered levels of MYH7B would impact
the developing heart is not understood, but this finding underscores the regulatory
complexity that likely underlies the involvement of all the CELF members.
171 In Chapter 6, we also describe the looping defects consequent to Celf1 knockdown. The CELF1 targets that underlie cardiac looping remain to be found. In zebrafish, dmrt2a was identified as one such target (Matsui et al., 2012), whose levels are normally kept low by celf1-mediated transcript degradation. Upon celf1 overexpression, defects of symmetry and somitogenesis ensue. However, the authors indicate that embryos injected with celf1-resistant dmrt2a exhibit the same defects, albeit at lower numbers. This suggests that additional celf1 targets play a role in this developmental process. Removal of celf1 also results in symmetry and laterality defects, as well as segmentation defects (Matsui et al., 2012) similar to those seen following Celf1 knockdown in frog embryos (Gautier-Courteille et al.,
2004). Taken together, these reports support the view that CELF1 is responsible for keeping at least some of its targets within a specific range of expression and that essential to this regulatory role is the careful maintenance of CELF1 itself within a narrow range of expression.
7.3. CELF1 in cardiac function of the embryonic heart
It is reasonable to speculate that the myofibrillar disorganization we observe in cultured cells and in developing hearts following CELF1 knockdown will result in functional deficits. In culture, primary cardiomyocytes continue to contract despite robust knockdown of CELF1 and profound myofibrillar disorganization. Embryonic hearts continue to beat even after Celf1 knockdown and despite myofibrillar disorganization and looping defects, although the appearance of ventral edema suggests some cardiac dysfunction. Cardiomyocytes also make up structural
172 components of the heart, such as septa and trabeculae, which are involved in the
proper flow of blood through the cardiac chambers. Defects in the formation or
remodeling of these structures may also impact the functional efficiency of the
heart, even in the absence of overt differences in contractile properties. Future
studies will evaluate the functional sequelae of Celf1 depletion in the embryonic
heart. Extensive analysis of the MHC-CELFΔ mice suggests that functional repression of CELF1 should lead to robust cardiac dysfunction (Ladd et al., 2005b).
However, in these mice, the nuclear activity of all myocardial CELF proteins is repressed, and this repression only begins after birth, making these mice an inappropriate system for studying the embryonic heart. Studies of mice in which
CELF1 is systemically inactivated have investigated neither the structure nor the function of the heart and, in fact, have largely focused only on the fertility defect observed in these animals (Cibois et al., 2012; Kress et al., 2007). These mice are an attractive model system for future studies. Other model organisms can also be used to answer this question. MOs can be microinjected into both frogs and zebrafish, followed by measurement of functional parameters.
7.4. Future directions
Techniques have been reported for the study of small hearts in some organisms using transmitted light video-microscopy (Fink et al., 2009; Ocorr et al.,
2009). The use of optical coherence tomography (OCT) for the non-invasive imaging
of live embryos in real time has recently begun to receive much attention (Garita et
al., 2011; Hoage et al., 2012; Manner et al., 2008). We have begun to utilize OCT for the analysis of cardiac morphology, as well as function, in frog embryos (not
173 shown). In future studies, we hope to use this approach to investigate the functional consequences of perturbing CELF1-mediated regulatory programs in the developing heart.
The ability to repress only the nuclear activity of Celf1 within the developing frog embryo will help dissect the contributions of nuclear and cytoplasmic roles in the myofibrillar disorganization we observe. In this way, the early role of Celf1 in the degradation of maternal transcripts during the mid-blastula transition can be circumvented. The dominant negative NLSCELFΔ will be microinjected into frog
embryos and cardiac structure and function will be evaluated. Cardiac endogenous
targets of Celf1-mediated alternative splicing regulation must be identified in order
to verify that the microinjected NLSCELFΔ is in fact repressing Celf1 activity in vivo,
as injection of a plasmid-bourn mini-gene reporter was found to result in
undetectable levels of transcript in stage 35-36 whole embryo extracts (not shown).
Finally, the investigation of Celf1-mediated functions in the developing heart can be
further refined by microinjection of Celf1, Celf1-MOs, or NLSCELFΔ only into
blastomeres fated to give rise primarily to the heart. In this way, neighboring tissues
will be spared.
The effect of CELF1 knockdown was described in isolated cardiomyocytes
and in developing hearts. Overexpression of transgenic CELF1 in muscle cells in the
mouse results in a muscular dystrophy phenotype. In skeletal muscle cells, in which
exit from the cell cycle is necessary for differentiation, over-expression of CELF1
results in enhanced proliferation, stalled skeletal muscle differentiation, and
disorganization of muscle fibers (Timchenko et al., 2004). A role for CELF1 in the
174 organization of the myofibril has yet to be investigated in skeletal muscle cells. We have preliminarily used immunofluorescence to evaluate myofibrillar organization in skeletal muscle in frog embryos injected with Celf1-MOs. Our findings suggest that knockdown of Celf1 results in disorganization of the muscle fibers without affecting the organization of the myofibrils (not shown). While these studies should be repeated, these preliminary observations suggest that the involvement of Celf1 in myofibrillar organization may be unique to cardiac muscle cells. Furthermore, evaluation of the effect of knockdown of Celf1 in the developing embryo on proliferation of cardiomyocytes (and skeletal muscle cells) can be easily carried out by injection of EdU into live embryos. The assessment will provide a better understanding of differences between cardiac and skeletal muscle cells, but will also indicate if the proliferation phenotype we observe in cultured cardiomyocytes
(Chapter 6) is also found in the developing heart. For instance, signals from other components in the developing tissue may supersede a role for CELF1 in this process in the context of the developing embryo.
The trans-regulation of CELF proteins by their paralogs remains largely unknown. CELF1 was shown to bind to the 3’ UTR of its own transcript by HITS-
CLIP and was shown to be able to both bind and regulate degradation of Celf2 in
C2C12 cultured myoblasts (Masuda et al., 2012). In an in vitro assay, CELF1 was shown to be able to bind to its own transcript, as well as to CELF2 (albeit to a lesser degree)(Lu et al., 1999). In fact, one target of CELF1 binding identified by CLIP analysis in our studies (Chapter 5) was CELF2 (Table S2). In an in vitro assay,
CELF2 was likewise shown to be able to bind both to its own transcript and to
175 CELF1 (Lu et al., 1999), and has also been proposed to regulate its own levels or function by regulating its splicing in a human embryonic kidney cell line, HEK293
(Dembowski and Grabowski, 2009). It also remains unclear to what degree CELF proteins may share targets. The great diversity of isoforms and subcellular localization of even just CELF1 and CELF2 (described in Chapter 4) suggests that these proteins are not merely redundant. The knockdown of CELF1 in cultured cells and in the developing embryo will be used to evaluate the role of CELF1 in the regulation of other CELF proteins, such as CELF2. Furthermore, the frog embryo is an ideal system for the over-expression of CELF1 or CELF2, and the evaluation of the effects of this manipulation both on these proteins (and transcripts) themselves and on candidate targets believed to be either shared or unique.
176 Appendix
Table S1. CELF1 CLIP tags obtained from embryonic day 8 chicken heart. (Appended as a digital file)
Table S2. The subset of intronic CELF1 CLIP tags that fall within annotated genes. (Appended as a digital file)
177 Supplemental Script 1 #! /usr/bin/perl use strict; use warnings;
#Written by Yotam Blech-Hermoni #Program in Cell Biology, Case Western Reserve University, Cleveland, OH #Laboratory of Dr. Andrea Ladd, PhD #Department of Cellular and Molecular Medicine, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH #Date: 10/20/2014 #Purpose: This program will identify sequence tags generated from a CLIP assay, followed by cloning. #Note 1: The input files used were *.seq files generated by an ABI GA sequencer. In order to facilitate the #batch file name generation, the .seq extension should be excluded when entering the filename within the program #(the program will append the extension on its own). For us, each filename consisted of the three-digit number #of the clone it represented. The filenames in the batch MUST be sequential and uninterrupted for this script. #Note 2: This script was written in order to solve two problems: 1. linkers often concatenated #during the process and 2. multiple tags were cloned into each plasmid. #Note 3: This script will generate one output file containing a list of tags from the range of input files entered. #Note 4: The linker sequences can be defined below.
print "\nThis program will find tags within sequence files\n\n";
#Prompt for first and last file names (numbers) print "What is the number of the first file in the batch\?"; $filenumber_first =
#Setup loop counter for batch of all files for ($i = $filenumber_first; $i < $filenumber_last; ++$i){ $filenumber = $i; $filename = $filenumber."\.seq";
#Define input template $original_template = extract_DNA($filename); $old_template = $original_template."55"; $template1 = $old_template; #Save copy for output file 178
#Define core linker sequences #If you are using different adapters, you should enter them here $L1_forward = "GAGGAC"; $L1_reverse = "CATCGT"; $L2_forward = "TCAGTC"; $L2_reverse = "GAAGTG";
#Substitute linker sequence with number designation $old_template =~ s/$L1_forward/1/g; $old_template =~ s/$L1_reverse/4/g; $old_template =~ s/$L2_forward/2/g; $old_template =~ s/$L2_reverse/3/g; $original_length = length $old_template; $template2 = $old_template; #Save copy for output file
#Define templates @old_template = split ('',$old_template); $new_template = ""; @new_template = split ('', $new_template);
#Define first four fragments (linker sequences) in output array @fragments = ""; $fragments[0] = "L1_forward\(1\) \= $L1_forward"; $fragments[1] = "L1_reverse\(2\) \= $L1_reverse"; $fragments[2] = "L2_forward\(3\) \= $L2_forward"; $fragments[3] = "L2_reverse\(4\) \= $L2_reverse";
#Initialize counter $j = 0;
#Carry out until there is no more old template left while ($j<$original_length){ do{ #Remove value from start of old template $popper = shift @old_template; $old_template = join ('', @old_template);
#Add value to end of new template push (@new_template, $popper); $new_template = join ('', @new_template); ++$j; }until ($new_template =~ /[1-5].*[1-5]/); $fragment = $&;
#Output length of tag to file 179 print OUTPUTFILE $fragments[$k+4]."\n"; print OUTPUTFILE "Length: ".$longness."nt\n\n"; } @fragments = ""; close(OUTPUTFILE); } exit; #------##SUBROUTINES##------sub extract_DNA{ my ($filename) = @_; open (FILENAME, $filename); my (@DNA) =
sub trim1{ my ($fragment) = @_; my (@tail_of_1) = ('G','A','T','G','C','G','G'); my ($b) = 0; do{ my ($end_of_1) = "1"."$tail_of_1[$b]"; if ($fragment =~ /$end_of_1/){ my ($motif) = $&; $fragment =~ s/$motif/1/; ++$b; } }while ($b < 7); return $fragment; }
sub trim2{ my ($fragment) = @_; my (@tail_of_2) = ('G','T','G'); my ($d) = 2; do{ my ($end_of_2) = "$tail_of_2[$d]"."2"; if ($fragment =~ /$end_of_2/){ my ($motif) = $&; $fragment =~ s/$motif/2/; --$d; } 180 }while ($d > -1); return $fragment; }
sub trim3{ my ($fragment) = @_; my (@tail_of_3) = ('A','C','T','G','A','C','A','C'); my ($p) = 0; do{ my ($end_of_3) = "3"."$tail_of_3[$p]"; if ($fragment =~ /$end_of_3/){ my ($motif) = $&; $fragment =~ s/$motif/3/; ++$p; } }while ($p < 8); return $fragment; }
sub trim4{ my ($fragment) = @_; my (@tail_of_4) = ('C','C','G'); my ($q) = 2; do{ my ($end_of_4) = "$tail_of_4[$q]"."4"; if ($fragment =~ /$end_of_4/){ my ($motif) = $&; $fragment =~ s/$motif/4/; --$q; } }while ($q > -1); return $fragment; }
181 Supplemental Script 2 #! /usr/bin/perl use strict; use warnings;
#Program name: super_tagalyzer.pl #Written by Yotam Blech-Hermoni #Program in Cell Biology, Case Western Reserve University, Cleveland, OH #Laboratory of Dr. Andrea Ladd, PhD #Department of Cellular and Molecular Medicine, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH #Date: 10/20/2014 #Purpose: This program will take [CLIP] tags from a file, generate a list of all possible motifs (n-mers; for 0 my @motif_tallies = (); ##Output array my $window = 0; ##Moving window of tag to be compared to motifs my $tag2 = 0; ##Tag to be analyzed my @tags = (); ##List-array of tags my @motifs = (); ##List-array of motifs my $infile = $ARGV[0]; chomp $infile; ##Open file open (TAGS, "$infile"); ##Put tag list into array: ##Take items in file - sequentially - as input while (my $tag1 = ##mer_number: ##The program will generate motifs between 1 and 6 nucleotides long. my $motif_length = $ARGV[1]; 182 if ($motif_length < 7){ ##Call motif generator (length < 7) gen_mod($motif_length); ##See subroutine below for attribution information! open (MOTIFS, "motifs"); ##Clean up motif list, and put motifs into array while (my $motif1 = ##For each tag... for (my $i = 0; $i < @tags; ++$i){ $tag2 = $tags[$i]; ##For as long as the tag lasts... for (my $n = 0; $n < length $tag2; ++$n){ ##Create a moving window $window = substr($tag2, $n, $motif_length); ##Compare each motif to the moving window for (my $j = 0; $j < @motifs; ++$j){ ##Pick a motif to compare to moving window my $motif = $motifs[$j]; if ($motif eq $window){ $motif_tallies[$j] += 1; }else {$motif_tallies[$j] += 0}; } } } print OUTFILE "@motif_tallies\n"; close OUTFILE; @motifs = (); @motif_tallies = (); }else{ for (my $motif_length = 1; $motif_length < 7; ++$motif_length){ gen_mod($motif_length); ##See subroutine below for attribution information! 183 open (MOTIFS, "motifs"); ##Clean up motif list, and put motifs into array while (my $motif1 = ##For each tag... for (my $i = 0; $i < @tags; ++$i){ $tag2 = $tags[$i]; ##For as long as the tag lasts... for (my $n = 0; $n < length $tag2; ++$n){ ##Create a moving window $window = substr($tag2, $n, $motif_length); ##Compare each motif to the moving window for (my $j = 0; $j < @motifs; ++$j){ ##Pick a motif to compare to moving window my $motif = $motifs[$j]; if ($motif eq $window){ $motif_tallies[$j] += 1; }else {$motif_tallies[$j] += 0}; } } } print OUTFILE "@motif_tallies\n"; close OUTFILE; @motifs = (); @motif_tallies = (); } } exit; #------##SUBROUTINES##------ sub gen_mod{ # MODIFIED FROM: 184 # 6gen.pl - generate all possible password # combinations up to 6 characters # # 2008 - Mike Golvach - [email protected] # # Creative Commons Attribution-Noncommercial-Share Alike 3.0 United States License # my ($length) = @_; if ( $length < 0 ) { print "\nUsage: $0 password_length\n"; print "Only 6 characters please. This is\n"; print "going to take a long time as it is!\n"; exit(1); } my $pass_length = $length; @4 = qw(A C G T); if ( $pass_length < 1 || $pass_length > 6 ) { print "Usage: $0 password_length\n"; print "Only 6 characters please. This is\n"; print "going to take a long time as it is!\n"; } my $c = my $d = my $e = my $f = my $g = my $h = 4; open MOTIFS, ">>motifs"; my $i = 0; if ( $pass_length == 6 ) { for ($c=0;$c<4;$c++) { for ($d=0;$d<4;$d++) { for ($e=0;$e<4;$e++) { for ($f=0;$f<4;$f++) { for ($g=0;$g<4;$g++) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s%s%s%s%s%s\n", $4[$c], $4[$d], $4[$e], $4[$f], $4[$g], $4[$h]); ++$i; } } } } } } } elsif ( $pass_length == 5 ) { for ($d=0;$d<4;$d++) { for ($e=0;$e<4;$e++) { for ($f=0;$f<4;$f++) { for ($g=0;$g<4;$g++) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s%s%s%s%s\n", $4[$d], $4[$e], $4[$f], $4[$g], $4[$h]); } } } } } } elsif ( $pass_length == 4 ) { for ($e=0;$e<4;$e++) { for ($f=0;$f<4;$f++) { for ($g=0;$g<4;$g++) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s%s%s%s\n", $4[$e], $4[$f], $4[$g], $4[$h]); } } } } } elsif ( $pass_length == 3 ) { for ($f=0;$f<4;$f++) { for ($g=0;$g<4;$g++) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s%s%s\n", $4[$f], $4[$g], $4[$h]); } } } 185 } elsif ( $pass_length == 2 ) { for ($g=0;$g<4;$g++) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s%s\n", $4[$g], $4[$h]); } } } elsif ( $pass_length == 1 ) { for ($h=0;$h<4;$h++) { printf MOTIFS ("%s\n", $4[$h]); } } close MOTIFS; } 186 Supplemental Script 3 #! /usr/bin/perl use strict; use warnings; #Program name: tag_mapper.pl #Written by Yotam Blech-Hermoni #Program in Cell Biology, Department of Molecular Biology and Microbiology, Case Western Researve University School of Medicine, Cleveland, OH #Laboratory of Dr. Andrea Ladd, PhD #Department of Cellular and Molecular Medicine, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH #Date: 10/20/14 #Purpose: Takes list of intronic tags as input. Takes list of chicken introns and compares tags to tag list to find distance of tags from intron ends. #Note 1: The unRandom chromosomes are not supported. #Note 2: Make sure there are no *.tmp files in the folder in which this script is run, as the script includes lines that will erase them. #Note 3: Files and file names that must be created or modified: #In &tag_file_generator: tag_list is the list of tags; consists of 6 columns: #C1: tag id #C2: chromosome (number) #C3: start #C4: end #C5: length #C6: strand #In &intron_file_generator : introns_galGal3 is the intron input file; consists of 6 columns (THIS FILE MUST BE PRE-SORTED FOR C1 AND C2): #The script is designed for input from UCSC. Input from other databases may require adjustments to coordinates and/or chromosome annotation. #C1: chromosome #C2: start #C3: end #C4: description #C6: strand #Output file: tag_mapper_gg_output.txt my $filename = $ARGV[0]; chomp $filename; ##generage *.tmp files and list-arrays of tag attributes &tag_file_generator; my @tag_chrs = &tags_chr; ##call subroutine to generate array of tag chromosomes my @tag_starts = &tags_start; ##call subroutine to generate array of tag starts my @tag_ends = &tags_end; ##call subroutine to generate array of tag ends my @tag_strands = &tags_strand;##call subroutine to generate array of tag strands 187 ##create array of reference chromosomes my @chromosomes = ('chr1', 'chr2', 'chr3', 'chr4', 'chr5', 'chr6', 'chr7', 'chr8', 'chr9', 'chr10', 'chr11', 'chr12', 'chr13', 'chr14', 'chr15', 'chr16', 'chr17', 'chr18', 'chr19', 'chr20', 'chr21', 'chr22', 'chr23', 'chr24', 'chr26', 'chr27', 'chr28', 'chr32', 'chrW', 'chrZ', 'chrM'); ##initialize counter my $i = 0; ##print header line to output file open (OUTPUT, ">>tag_mapper_gg_output.txt"); print OUTPUT "row_number\t"; print OUTPUT "chr\t"; print OUTPUT "tag_start\t"; print OUTPUT "tag_end\t"; print OUTPUT "direction\t"; print OUTPUT "intron_description\t"; print OUTPUT "intron_start\t"; print OUTPUT "intron_end\t"; print OUTPUT "intron_length\t"; print OUTPUT "upstream_distance\t"; print OUTPUT "downstream_distance\n"; #main loops comparing tag chromosome list to reference chromosome list and mapping tags to matched chromosomes foreach my $tag_chr (@tag_chrs){ ##take each tag chromosome foreach my $chr (@chromosomes){ ##take each reference chromosome if ($tag_chr eq $chr){ ##if reference chromosome matches tag… my $row_number = $i + 1; ##extract tag attributes my $tag_start = $tag_starts[$i]; my $tag_end = $tag_ends[$i]; my $tag_strand = $tag_strands[$i]; $tag_strand =~ tr/-+/rf/; system "rm \*\.tmp"; ##generate file of all introns in matched chromosome &intron_file_generator(\$chr); ##generate list-arrays of all intron attributes in matched chromosome my @intron_starts = &introns_start; my @intron_ends = &introns_end; my @descriptions = &descriptions; my @intron_strands = &introns_strand; 188 ##call subroutine that calculates the intron in which the tag resides my $intron = &calc_intron($tag_start, $tag_end, \@intron_starts, \@intron_ends); if ($intron < 0){ print OUTPUT "Tag is not within an annotated intron\n"; }else { ##extract intron attributes my $intron_start = $intron_starts[$intron]; my $intron_end = $intron_ends[$intron]; my $intron_length = $intron_end - $intron_start; my $intron_description = $descriptions[$intron]; my $intron_strand = $intron_strands[$intron]; $intron_strand =~ tr/-+/rf/; ##calculate distance between tag and intron ends, with attention to the direction of the tag ##if the tag and intron are on opposite strands, output message to that effect my $up_distance = ''; my $down_distance = ''; if ($tag_strand eq "f" && $intron_strand eq "r"){ $up_distance = "Tag is on forward strand\n"; $down_distance = "Intron is on reverse strand\n"; }elsif ($tag_strand eq "r" && $intron_strand eq "f"){ $up_distance = "Tag is on reverse strand\n"; $down_distance = "Intron is on forward strand\n"; }elsif ($tag_strand eq "f" && $intron_strand eq "f"){ $up_distance = $tag_start - $intron_start; $down_distance = $intron_end - $tag_end; }elsif ($tag_strand eq "r" && $intron_strand eq "r"){ $up_distance = $intron_end - $tag_end; $down_distance = $tag_start - $intron_start; } print OUTPUT "$row_number\t"; print OUTPUT "$chr\t"; print OUTPUT "$tag_start\t"; print OUTPUT "$tag_end\t"; print OUTPUT "$tag_strand\t"; print OUTPUT "$intron_description\t"; print OUTPUT "$intron_start\t"; print OUTPUT "$intron_end\t"; print OUTPUT "$intron_length\t"; 189 print OUTPUT "$up_distance\t"; print OUTPUT "$down_distance\n"; } }else { next; } } ++$i; } close (OUTPUT); system "rm \*\.tmp"; exit; #------##SUBROUTINES##------sub tag_file_generator{ ##from tag_list file, extract columns and create *.tmp files for each tag attribute system "cat $filename \| cut \-f1 \>tags\_id\.tmp\n"; system "cat $filename \| cut \-f2 \>tags\_chr\.tmp\n"; system "cat $filename \| cut \-f3 \>tags\_start\.tmp\n"; system "cat $filename \| cut \-f4 \>tags\_end\.tmp\n"; system "cat $filename \| cut \-f5 \>tags\_length\.tmp\n"; system "cat $filename \| cut \-f6 \>tags\_strand\.tmp\n"; } sub tags_chr{ ##transfer tag chromosome list from *.tmp file into array open (IN, "tags_chr.tmp"); my $n = 0; my @tag_chrs = (); while ( sub tags_start{ ##transfer tag starts list from *.tmp file into array open (IN, "tags_start.tmp"); my $n = 0; my @tag_starts = (); while ( sub tags_end{ ##transfer tag ends list from *.tmp file into array open (IN, "tags_end.tmp"); my $n = 0; my @tag_ends = (); while ( sub tags_strand{ ##transfer tag strands list from *.tmp file into array open (IN, "tags_strand.tmp"); my $n = 0; my @tag_strands = (); while ( sub descriptions{ ##transfer intron descriptions list from *.tmp file into array open (IN, "descriptions_by_chromosome.tmp"); my $n = 0; my @descriptions = (); while ( 193 References Adamcova, M., Pelouch, V., 1999. Isoforms of troponin in normal and diseased myocardium. Physiol Res 48, 235-247. Adereth, Y., Dammai, V., Kose, N., Li, R., Hsu, T., 2005. RNA-dependent integrin alpha3 protein localization regulated by the Muscleblind-like protein MLP1. Nat Cell Biol 7, 1240-1247. Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z., Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25, 3389-3402. Amato, M.A., Boy, S., Arnault, E., Girard, M., Della Puppa, A., Sharif, A., Perron, M., 2005. Comparison of the expression patterns of five neural RNA binding proteins in the Xenopus retina. J Comp Neurol 481, 331-339. Amaya, E., Offield, M.F., Grainger, R.M., 1998. Frog genetics: Xenopus tropicalis jumps into the future. Trends Genet 14, 253-255. Ames, E.G., Lawson, M.J., Mackey, A.J., Holmes, J.W., 2013. Sequencing of mRNA identifies re- expression of fetal splice variants in cardiac hypertrophy. J Mol Cell Cardiol 62C, 99- 107. Amiel, J., de Pontual, L., Henrion-Caude, A., 2012. miRNA, development and disease. Adv Genet 80, 1-36. Anant, S., Henderson, J., Mukhopadhyay, D., Navaratnam, N., Kennedy, S., Min, J., Davidson, N., 2001. Novel role for RNA-binding protein CUGBP2 in mammalian RNA editing. J Biol Chem 276, 47338-47351. Anderson, J.T., Wilson, S.M., Datar, K.V., Swanson, M.S., 1993. NAB2: a yeast nuclear polyadenylated RNA-binding protein essential for cell viability. Mol Cell Biol 13, 2730-2741. Arthur, H.M., Bamforth, S.D., 2011. TGFbeta signaling and congenital heart disease: Insights from mouse studies. Birth Defects Res A Clin Mol Teratol 91, 423-434. Ashe M, B.D., 2005. Development of the Vertebrate Heart. Humana Press Inc. Ast, G., 2004. How did alternative splicing evolve? Nat Rev Genet 5, 773-782. Audic, Y., Omilli, F., Osborne, H.B., 1998. Embryo deadenylation element-dependent deadenylation is enhanced by a cis element containing AUU repeats. Mol Cell Biol 18, 6879-6884. Baehrecke, E.H., 1997. who encodes a KH RNA binding protein that functions in muscle development. Development 124, 1323-1332. Baldwin, B.R., Timchenko, N.A., Zahnow, C.A., 2004. Epidermal growth factor receptor stimulation activates the RNA binding protein CUG-BP1 and increases expression of C/EBPbeta-LIP in mammary epithelial cells. Mol Cell Biol 24, 3682-3691. Balza, R.O., Jr., Misra, R.P., 2006. Role of the serum response factor in regulating contractile apparatus gene expression and sarcomeric integrity in cardiomyocytes. J Biol Chem 281, 6498-6510. Barash, Y., Calarco, J.A., Gao, W., Pan, Q., Wang, X., Shai, O., Blencowe, B.J., Frey, B.J., 2010. Deciphering the splicing code. Nature 465, 53-59. 194 Barbosa-Morais, N.L., Carmo-Fonseca, M., Aparicio, S., 2006. Systematic genome-wide annotation of spliceosomal proteins reveals differential gene family expansion. Genome Res 16, 66-77. Barreau, C., Paillard, L., Mereau, A., Osborne, H.B., 2006. Mammalian CELF/Bruno-like RNA- binding proteins: molecular characteristics and biological functions. Biochimie 88, 515-525. Barron, V.A., Zhu, H., Hinman, M.N., Ladd, A.N., Lou, H., 2010. The neurofibromatosis type I pre-mRNA is a novel target of CELF protein-mediated splicing regulation. Nuc Acids Res 38, 253-264. Beisang, D., Rattenbacher, B., Vlasova-St Louis, I.A., Bohjanen, P.R., 2012. Regulation of CUG- binding protein 1 (CUGBP1) binding to target transcripts upon T cell activation. J Biol Chem 287, 950-960. Belaguli, N.S., Zhou, W., Trinh, T.H., Majesky, M.W., Schwartz, R.J., 1999. Dominant negative murine serum response factor: alternative splicing within the activation domain inhibits transactivation of serum response factor binding targets. Mol Cell Biol 19, 4582-4591. Bell, M.L., Buvoli, M., Leinwand, L.A., 2010. Uncoupling of expression of an intronic microRNA and its myosin host gene by exon skipping. Mol Cell Biol 30, 1937-1945. Berger, D.S., Ladd, A.N., 2012. Repression of nuclear CELF activity can rescue CELF- regulated alternative splicing defects in skeletal muscle models of myotonic dystrophy. PLoS Curr 4, RRN1305. Berger, D.S., Moyer, M., Kliment, G.M., van Lunteren, E., Ladd, A.N., 2011. Expression of a dominant negative CELF protein in vivo leads to altered muscle organization, fiber size, and subtype. PLoS One 6, e19274. Besse, F., Ephrussi, A., 2008. Translational control of localized mRNAs: restricting protein synthesis in space and time. Nat Rev Mol Cell Biol 9, 971-980. Bland, C.S., Wang, E.T., Vu, A., David, M.P., Castle, J.C., Johnson, J.M., Burge, C.B., Cooper, T.A., 2010. Global regulation of alternative splicing during myogenic differentiation. Nuc Acids Res 38, 7651-7664. Blech-Hermoni, Y., Stillwagon, S.J., Ladd, A.N., 2013. Diversity and conservation of CELF1 and CELF2 RNA and protein expression patterns during embryonic development. Dev Dyn 242, 767-777. Blitz, I.L., Andelfinger, G., Horb, M.E., 2006. Germ layers to organs: using Xenopus to study "later" development. Semin Cell Dev Biol 17, 133-145. Bodmer, R., 1993. The gene tinman is required for specification of the heart and visceral muscles in Drosophila. Development 118, 719-729. Boutz, P.L., Stoilov, P., Li, Q., Lin, C.H., Chawla, G., Ostrow, K., Shiue, L., Ares, M., Jr., Black, D.L., 2007. A post-transcriptional regulatory switch in polypyrimidine tract-binding proteins reprograms alternative splicing in developing neurons. Genes Dev 21, 1636-1652. Bowers, S.L., Banerjee, I., Baudino, T.A., 2010. The extracellular matrix: at the center of it all. J Mol Cell Cardiol 48, 474-482. Bowes, J.B., Snyder, K.A., Segerdell, E., Gibb, R., Jarabek, C., Noumen, E., Pollet, N., Vize, P.D., 2008. Xenbase: a Xenopus biology and genomics resource. Nuc Acids Res 36, D761- 767. 195 Bradford, Y., Conlin, T., Dunn, N., Fashena, D., Frazer, K., Howe, D.G., Knight, J., Mani, P., Martin, R., Moxon, S.A., Paddock, H., Pich, C., Ramachandran, S., Ruef, B.J., Ruzicka, L., Bauer Schaper, H., Schaper, K., Shao, X., Singer, A., Sprague, J., Sprunger, B., Van Slyke, C., Westerfield, M., 2011. ZFIN: enhancements and updates to the Zebrafish Model Organism Database. Nuc Acids Res 39, D822-829. Brand, T., 2003. Heart development: molecular insights into cardiac specification and early morphogenesis. Dev Biol 258, 1-19. Brandenberger, R., Wei, H., Zhang, S., Lei, S., Murage, J., Fisk, G.J., Li, Y., Xu, C., Fang, R., Guegler, K., Rao, M.S., Mandalam, R., Lebkowski, J., Stanton, L.W., 2004. Transcriptome characterization elucidates signaling networks that control human ES cell growth and differentiation. Nature biotechnology 22, 707-716. Brauch, K.M., Karst, M.L., Herron, K.J., de Andrade, M., Pellikka, P.A., Rodeheffer, R.J., Michels, V.V., Olson, T.M., 2009. Mutations in ribonucleic acid binding protein gene cause familial dilated cardiomyopathy. J Am Coll Cardiol 54, 930-941. Brimacombe, K.R., Ladd, A.N., 2007. Cloning and embryonic expression patterns of the chicken CELF family. Dev Dyn 236, 2216-2224. Brudno, M., Gelfand, M., Spengler, S., Zorn, M., Dubchak, I., Conboy, J., 2001. Computational analysis of candidate intron regulatory elements for tissue-specific alternative pre- mRNA splicing. Nuc Acids Res 29, 2338-2348. Burd, C., Dreyfuss, G., 1994. Conserved structures and diversity of functions of RNA-binding proteins. Science 265, 615-621. Burt, D.W., Carre, W., Fell, M., Law, A.S., Antin, P.B., Maglott, D.R., Weber, J.A., Schmidt, C.J., Burgess, S.C., McCarthy, F.M., 2009. The Chicken Gene Nomenclature Committee report. BMC Genomics 10 Suppl 2, S5. Castle, J.C., Zhang, C., Shah, J.K., Kulkarni, A.V., Kalsotra, A., Cooper, T.A., Johnson, J.M., 2008. Expression of 24,426 human alternative splicing events and predicted cis regulation in 48 tissues and cell lines. Nat Genet 40, 1416-1425. Chang, E.T., Donahue, J.M., Xiao, L., Cui, Y., Rao, J.N., Turner, D.J., Twaddell, W.S., Wang, J.Y., Battafarano, R.J., 2012. The RNA-binding protein CUG-BP1 increases survivin expression in oesophageal cancer cells through enhanced mRNA stability. Biochem J 446, 113-123. Chapnik, E., Sasson, V., Blelloch, R., Hornstein, E., 2012. Dgcr8 controls neural crest cells survival in cardiovascular development. Dev Biol 362, 50-56. Charlet-B., N., Logan, P., Singh, G., Cooper, T., 2002a. Dynamic antagonism between ETR-3 and PTB regulates cell type-specific alternative splicing. Mol Cell 9, 649-658. Charlet-B., N., Savkur, R., Singh, G., Philips, A., Grice, E., Cooper, T., 2002b. Loss of the muscle-specific chloride channel in type I myotonic dystrophy lead to misregulated alternative splicing. Mol Cell 10, 45-53. Chateigner-Boutin, A.L., Small, I., 2011. Organellar RNA editing. Wiley Interdiscip Rev RNA 2, 493-506. Chen, H.W., Yu, S.L., Chen, W.J., Yang, P.C., Chien, C.T., Chou, H.Y., Li, H.N., Peck, K., Huang, C.H., Lin, F.Y., Chen, J.J., Lee, Y.T., 2004. Dynamic changes of gene expression profiles during postnatal development of the heart in mice. Heart 90, 927-934. Chen, J., Wang, D.Z., 2012. microRNAs in cardiovascular development. J Mol Cell Cardiol 52, 949-957. 196 Choi, D., Ito, T., Tsukahara, F., Hirai, M., Sakaki, Y., 1999. Developmentally regulated expression of mNapor encoding an apoptosis-induced ELAV-type RNA binding protein. Gene 237, 135-142. Choi, D.K., Yoo, K.W., Hong, S.K., Rhee, M., Sakaki, Y., Kim, C.H., 2003. Isolation and expression of Napor/CUG-BP2 in embryo development. Biochem Biophys Res Commun 305, 448-454. Cibois, M., Boulanger, G., Audic, Y., Paillard, L., Gautier-Courteille, C., 2012. Inactivation of the Celf1 gene that encodes an RNA-binding protein delays the first wave of spermatogenesis in mice. PLoS One 7, e46337. Cibois, M., Gautier-Courteille, C., Kodjabachian, L., Paillard, L., 2013. A gene regulation network controlled by Celf1 protein-rbpj mRNA interaction in Xenopus somite segmentation. Biol Open 2, 1078-1083. Cibois, M., Gautier-Courteille, C., Vallee, A., Paillard, L., 2010. A strategy to analyze the phenotypic consequences of inhibiting the association of an RNA-binding protein with a specific RNA. RNA 16, 10-15. Cloonan, N., Forrest, A.R., Kolle, G., Gardiner, B.B., Faulkner, G.J., Brown, M.K., Taylor, D.F., Steptoe, A.L., Wani, S., Bethel, G., Robertson, A.J., Perkins, A.C., Bruce, S.J., Lee, C.C., Ranade, S.S., Peckham, H.E., Manning, J.M., McKernan, K.J., Grimmond, S.M., 2008. Stem cell transcriptome profiling via massive-scale mRNA sequencing. Nat Methods 5, 613-619. Cooper, T.A., 2005. Use of minigene systems to dissect alternative splicing elements. Methods 37, 331-340. Danecek, P., Nellaker, C., McIntyre, R.E., Buendia-Buendia, J.E., Bumpstead, S., Ponting, C.P., Flint, J., Durbin, R., Keane, T.M., Adams, D.J., 2012. High levels of RNA-editing site conservation amongst 15 laboratory mouse strains. Genome biology 13, 26. Das, D., Clark, T.A., Schweitzer, A., Yamamoto, M., Marr, H., Arribere, J., Minovitsky, S., Poliakov, A., Dubchak, I., Blume, J.E., Conboy, J.G., 2007. A correlation with exon expression approach to identify cis-regulatory elements for tissue-specific alternative splicing. Nuc Acids Res 35, 4845-4857. Dasgupta, T., Ladd, A.N., 2012. The importance of CELF control: molecular and biological roles of the CUG-BP, Elav-like family of RNA-binding proteins. Wiley Interdiscip Rev RNA 3, 104-121. Dasgupta, T., Stillwagon, S.J., Ladd, A.N., 2013. Gene expression analyses implicate an alternative splicing program in regulating contractile gene expression and serum response factor activity in mice. PLoS One 8, e56590. Daughters, R.S., Tuttle, D.L., Gao, W., Ikeda, Y., Moseley, M.L., Ebner, T.J., Swanson, M.S., Ranum, L.P., 2009. RNA gain-of-function in spinocerebellar ataxia type 8. PLoS Genet 5, e1000600. De Craene, B., Berx, G., 2013. Regulatory networks defining EMT during cancer initiation and progression. Nat Rev Cancer 13, 97-110. DeHaan, R., 1968. Emergence of form and function in the embryonic heart. Dev Biol Suppl. 2, 208-250. DeHaan, R.L., 1969. Electrical recording from embryonic heart cells isolated in tissue culture. Experientia Suppl 15, 174-175. DeHaan, R.L., 1970. The potassium-sensitivity of isolated embryonic heart cells increases with development. Dev Biol 23, 226-240. 197 DeLaughter, D.M., Saint-Jean, L., Baldwin, H.S., Barnett, J.V., 2011. What chick and mouse models have taught us about the role of the endocardium in congenital heart disease. Birth Defects Res A Clin Mol Teratol 91, 511-525. Delaunay, J., Le Mee, G., Ezzeddine, N., Labesse, G., Terzian, C., Capri, M., Ait-Ahmed, O., 2004. The Drosophila Bruno paralogue Bru-3 specifically binds the EDEN translational repression element. Nuc Acids Res 32, 3070-3082. Dembowski, J.A., Grabowski, P.J., 2009. The CUGBP2 splicing factor regulates an ensemble of branchpoints from perimeter binding sites with implications for autoregulation. PLoS Genet 5, e1000595. Deutsch, M., Long, M., 1999. Intron-exon structures of eukaryotic model organisms. Nuc Acids Res 27, 3219-3228. Dev, A., Nayernia, K., Meins, M., Adham, I., Lacone, F., Engel, W., 2007. Mice deficient for RNA-binding protein brunol1 show reduction of spermatogenesis but are fertile. Mol Reprod Dev 74, 1456-1464. Diez-Roux, G., Banfi, S., Sultan, M., Geffers, L., Anand, S., Rozado, D., Magen, A., Canidio, E., Pagani, M., Peluso, I., Lin-Marq, N., Koch, M., Bilio, M., Cantiello, I., Verde, R., De Masi, C., Bianchi, S.A., Cicchini, J., Perroud, E., Mehmeti, S., Dagand, E., Schrinner, S., Nurnberger, A., Schmidt, K., Metz, K., Zwingmann, C., Brieske, N., Springer, C., Hernandez, A.M., Herzog, S., Grabbe, F., Sieverding, C., Fischer, B., Schrader, K., Brockmeyer, M., Dettmer, S., Helbig, C., Alunni, V., Battaini, M.A., Mura, C., Henrichsen, C.N., Garcia-Lopez, R., Echevarria, D., Puelles, E., Garcia-Calero, E., Kruse, S., Uhr, M., Kauck, C., Feng, G., Milyaev, N., Ong, C.K., Kumar, L., Lam, M., Semple, C.A., Gyenesei, A., Mundlos, S., Radelof, U., Lehrach, H., Sarmientos, P., Reymond, A., Davidson, D.R., Dolle, P., Antonarakis, S.E., Yaspo, M.L., Martinez, S., Baldock, R.A., Eichele, G., Ballabio, A., 2011. A high-resolution anatomical atlas of the transcriptome in the mouse embryo. PLoS Biol 9, e1000582. Ding, J.-H., Xu, X., Yang, D., Chu, P.-H., Dalton, N., Ye, Z., Yeakley, J., Cheng, H., Xiao, R.-P., Ross, J., Chen, J., Fu, X.-D., 2004. Dilated cardiomyopathy caused by tissue-specific ablation of SC35 in heart. EMBO J 23, 885-896. Dittmar, K.A., Jiang, P., Park, J.W., Amirikian, K., Wan, J., Shen, S., Xing, Y., Carstens, R.P., 2012. Genome-wide determination of a broad ESRP-regulated posttranscriptional network by high-throughput sequencing. Mol Cell Biol 32, 1468-1482. Dougherty, J.D., Maloney, S.E., Wozniak, D.F., Rieger, M.A., Sonnenblick, L., Coppola, G., Mahieu, N.G., Zhang, J., Cai, J., Patti, G.J., Abrahams, B.S., Geschwind, D.H., Heintz, N., 2013. The disruption of Celf6, a gene identified by translational profiling of serotonergic neurons, results in autism-related behaviors. J Neurosci 33, 2732- 2753. Dunwoodie, S.L., 2007. Combinatorial signaling in the heart orchestrates cardiac induction, lineage specification and chamber formation. Semin Cell Dev Biol 18, 54-66. Edmondson, D.G., Lyons, G.E., Martin, J.F., Olson, E.N., 1994. Mef2 gene expression marks the cardiac and skeletal muscle lineages during mouse embryogenesis. Development 120, 1251-1263. Edwards, J.M., Long, J., de Moor, C.H., Emsley, J., Searle, M.S., 2013. Structural insights into the targeting of mRNA GU-rich elements by the three RRMs of CELF1. Nuc Acids Res 41, 7153-7166. 198 Engels, B., van 't Padje, S., Blonden, L., Severijnen, L.A., Oostra, B.A., Willemsen, R., 2004. Characterization of Fxr1 in Danio rerio; a simple vertebrate model to study costamere development. J Exp Biol 207, 3329-3338. Eppig, J., 2006. Mouse Strain and Genetic Nomenclature: an Abbreviated Guide, in: Fox J, B.S., Davvison M, Newcomer C, Quimby F, Smith A (Ed.), The Mouse in Biomedical Research, 2nd ed. Academic Press, pp. 79-98. Esposito, T., Sampaolo, S., Limongelli, G., Varone, A., Formicola, D., Diodato, D., Farina, O., Napolitano, F., Pacileo, G., Gianfrancesco, F., Di Iorio, G., 2013. Digenic mutational inheritance of the integrin alpha 7 and the myosin heavy chain 7B genes causes congenital myopathy with left ventricular non-compact cardiomyopathy. Orphanet J Rare Dis 8, 91. Ewen, E.P., Snyder, C.M., Wilson, M., Desjardins, D., Naya, F.J., 2011. The Mef2A transcription factor coordinately regulates a costamere gene program in cardiac muscle. J Biol Chem 286, 29644-29653. Ezzeddine, N., Paillard, L., Capri, M., Maniey, D., Bassez, T., Ait-Ahmed, O., Osborne, H.B., 2002. EDEN-dependent translational repression of maternal mRNAs is conserved between Xenopus and Drosophila. Proceedings of the National Academy of Sciences of the United States of America 99, 257-262. Faber., P.D.N.a.J., 1967. Normal table of Xenopus laevis (Daudin) : a systematical and chronological survey of the development from the fertilized egg till the end of metamorphosis, 2nd ed. Hubrecht-Laboratorium (Embryologisch Instituut), Amsterdam. Fan, C., Chen, Q., Wang, Q.K., 2009. Functional role of transcriptional factor TBX5 in pre- mRNA splicing and Holt-Oram syndrome via association with SC35. The Journal of biological chemistry 284, 25653-25663. Fardaei, M., Larkin, K., Brook, J., Hamshere, M., 2001. In vivo co-localization of MBNL protein with DMPK expanded-repeat transcripts. Nuc Acids Res 29, 2766-2771. Fardaei, M., Rogers, M., Thorpe, H., Larkin, K., Hamshere, M., Harper, P., Brook, J., 2002. Three proteins, MBNL, MBLL and MBXL, co-localize in vivo with nuclear foci of expanded-repeat transcripts in DM1 and DM2 cells. Hum Mol Gen 11, 805-814. Faustino, N., Cooper, T., 2005. Identification of putative new splicing targets for ETR-3 using its SELEX sequences. Mol Cell Biol 25, 879-887. Feng, Y., Valley, M.T., Lazar, J., Yang, A.L., Bronson, R.T., Firestein, S., Coetzee, W.A., Manley, J.L., 2009. SRp38 regulates alternative splicing and is required for Ca(2+) handling in the embryonic heart. Dev Cell 16, 528-538. Fernandes, J.M., Kinghorn, J.R., Johnston, I.A., 2007. Characterization of two paralogous muscleblind-like genes from the tiger pufferfish (Takifugu rubripes). Comp Biochem Physiol B Biochem Mol Biol 146, 180-186. Fetka, I., Radeghieri, A., Bouwmeester, T., 2000. Expression of the RNA recognition motif- containing protein SEB-4 during Xenopus embryonic development. Mech Dev 94, 283-286. Fink, M., Callol-Massot, C., Chu, A., Ruiz-Lozano, P., Izpisua Belmonte, J.C., Giles, W., Bodmer, R., Ocorr, K., 2009. A new method for detection and quantification of heartbeat parameters in Drosophila, zebrafish, and embryonic mouse hearts. Biotechniques 46, 101-113. 199 Fomovsky, G.M., Thomopoulos, S., Holmes, J.W., 2010. Contribution of extracellular matrix to the mechanical properties of the heart. J Molec Cell Cardiol 48, 490-496. Fujimura, K., Kano, F., Murata, M., 2008. Dual localization of the RNA binding protein CUGBP-1 to stress granule and perinucleolar compartment. Exp Cell Res 314, 543- 553. Gallagher, T.L., Arribere, J.A., Geurts, P.A., Exner, C.R., McDonald, K.L., Dill, K.K., Marr, H.L., Adkar, S.S., Garnett, A.T., Amacher, S.L., Conboy, J.G., 2011. Rbfox-regulated alternative splicing is critical for zebrafish cardiac and skeletal muscle functions. Dev Biol 359, 251-261. Gao, F.B., Taylor, J.P., 2012. RNA-binding proteins in neurological disease. Brain Res 1462, 1-2. Garg, V., Yamagishi, C., Hu, T., Kathiriya, I.S., Yamagishi, H., Srivastava, D., 2001. Tbx1, a DiGeorge syndrome candidate gene, is regulated by sonic hedgehog during pharyngeal arch development. Dev Biol 235, 62-73. Garita, B., Jenkins, M.W., Han, M., Zhou, C., Vanauker, M., Rollins, A.M., Watanabe, M., Fujimoto, J.G., Linask, K.K., 2011. Blood flow dynamics of one cardiac cycle and relationship to mechanotransduction and trabeculation during heart looping. American journal of physiology. Am J Physiol Heart Circ Physiol 300, H879-891. Gates, D.P., Coonrod, L.A., Berglund, J.A., 2011. Autoregulated splicing of muscleblind-like 1 (MBNL1) Pre-mRNA. J Biol Chem 286, 34224-34233. Gautier-Courteille, C., Le Clainche, C., Barreau, C., Audic, Y., Graindorge, A., Maniey, D., Osborne, H.B., Paillard, L., 2004. EDEN-BP-dependent post-transcriptional regulation of gene expression in Xenopus somitic segmentation. Development 131, 6107-6117. Gerber, W.V., Vokes, S.A., Zearfoss, N.R., Krieg, P.A., 2002. A role for the RNA-binding protein, hermes, in the regulation of heart development. Dev Biol 247, 116-126. Gerber, W.V., Yatskievych, T.A., Antin, P.B., Correia, K.M., Conlon, R.A., Krieg, P.A., 1999. The RNA-binding protein gene, hermes, is expressed at high levels in the developing heart. Mech Dev 80, 77-86. Giardine, B., Riemer, C., Hardison, R.C., Burhans, R., Elnitski, L., Shah, P., Zhang, Y., Blankenberg, D., Albert, I., Taylor, J., Miller, W., Kent, W.J., Nekrutenko, A., 2005. Galaxy: a platform for interactive large-scale genome analysis. Genome Res 15, 1451-1455. Giudice, J., Xia, Z., Wang, E.T., Scavuzzo, M.A., Ward, A.J., Kalsotra, A., Wang, W., Wehrens, X.H., Burge, C.B., Li, W., Cooper, T.A., 2014. Alternative splicing regulates vesicular trafficking genes in cardiomyocytes during postnatal heart development. Nat Commun 5, 3603. Glaser, N.D., Lukyanenko, Y.O., Wang, Y., Wilson, G.M., Rogers, T.B., 2006. JNK activation decreases PP2A regulatory subunit B56alpha expression and mRNA stability and increases AUF1 expression in cardiomyocytes. Am J Physiol Heart Circ Physiol 291, H1183-1192. Glisovic, T., Bachorik, J.L., Yong, J., Dreyfuss, G., 2008. RNA-binding proteins and post- transcriptional gene regulation. FEBS letters 582, 1977-1986. Godt, R., Fogaca, R., Silva, I., Nosek, T., 1993. Contraction of developing avian heart muscle. Comp Biochem Physiol [A] 105, 213-218. 200 Goers, E.S., Purcell, J., Voelker, R.B., Gates, D.P., Berglund, J.A., 2010. MBNL1 binds GC motifs embedded in pyrimidines to regulate alternative splicing. Nuc Acids Res 38, 2467- 2484. Goo, Y.H., Cooper, T.A., 2009. CUGBP2 directly interacts with U2 17S snRNP components and promotes U2 snRNA binding to cardiac troponin T pre-mRNA. Nuc Acids Res 37, 4275-4286. Good, P., Chen, Q., Warner, S., Herring, D., 2000. A family of human RNA-binding proteins related to the Drosophila Bruno translational regulator. J Biol Chem 275, 28583- 28592. Gott, J.M., Emeson, R.B., 2000. Functions and mechanisms of RNA editing. Annu Rev Genet 34, 499-531. Gouble, A., Morello, D., 2000. Synchronous and regulated expression of two AU-binding proteins, AUF1 and HuR, throughout murine development. Oncogene 19, 5377- 5384. Graindorge, A., Le Tonqueze, O., Thuret, R., Pollet, N., Osborne, H.B., Audic, Y., 2008. Identification of CUG-BP1/EDEN-BP target mRNAs in Xenopus tropicalis. Nuc Acids Res 36, 1861-1870. Gray, K.A., Daugherty, L.C., Gordon, S.M., Seal, R.L., Wright, M.W., Bruford, E.A., 2013. Genenames.org: the HGNC resources in 2013. Nuc Acids Res 41, D545-552. Gregorio, C.C., Antin, P.B., 2000. To the heart of myofibril assembly. Trends Cell Biol 10, 355-362. Gripp, K.W., Hopkins, E., Johnston, J.J., Krause, C., Dobyns, W.B., Biesecker, L.G., 2011. Long- term survival in TARP syndrome and confirmation of RBM10 as the disease-causing gene. Am J Med Genet A 155A, 2516-2520. Gromak, N., Matlin, A., Cooper, T., Smith, C., 2003. Antagonistic regulation of α-actinin alternative splicing by CELF proteins and polypyrimidine tract binding protein. RNA 9, 443-456. Guo, W., Schafer, S., Greaser, M.L., Radke, M.H., Liss, M., Govindarajan, T., Maatz, H., Schulz, H., Li, S., Parrish, A.M., Dauksaite, V., Vakeel, P., Klaassen, S., Gerull, B., Thierfelder, L., Regitz-Zagrosek, V., Hacker, T.A., Saupe, K.W., Dec, G.W., Ellinor, P.T., MacRae, C.A., Spallek, B., Fischer, R., Perrot, A., Ozcelik, C., Saar, K., Hubner, N., Gotthardt, M., 2012. RBM20, a gene for hereditary cardiomyopathy, regulates titin splicing. Nat Med 18, 766-773. Halgren, C., Bache, I., Bak, M., Myatt, M.W., Anderson, C.M., Brondum-Nielsen, K., Tommerup, N., 2012. Haploinsufficiency of CELF4 at 18q12.2 is associated with developmental and behavioral disorders, seizures, eye manifestations, and obesity. Eur J Hum Genet. Hamburger, V., Hamilton, H.L., 1992. A series of normal stages in the development of the chick embryo. 1951. Dev Dyn 195, 231-272. Han, J., Cooper, T.A., 2005. Identification of CELF splicing activation and repression domains in vivo. Nuc Acids Res 33, 2769-2780. Harrell, M.D., Harbi, S., Hoffman, J.F., Zavadil, J., Coetzee, W.A., 2007. Large-scale analysis of ion channel gene expression in the mouse heart during perinatal development. Physiol Genomics 28, 273-283. Harvey, R., Rosenthal, N., 1999. Heart Development. Academic Press, In.c, San Diego, p. 530. Hertel, K.J., 2008. Combinatorial control of exon recognition. J Biol Chem 283, 1211-1215. 201 Ho, T., Charlet-B., N., Poulos, M., Singh, G., Swanson, M., Cooper, T., 2004. Muscleblind proteins regulate alternative splicing. EMBO J 23, 3103-3112. Ho, T.H., Bundman, D., Armstrong, D.L., Cooper, T.A., 2005. Transgenic mice expressing CUG-BP1 reproduce splicing mis-regulation observed in myotonic dystrophy. Hum Mol Genet 14, 1539-1547. Hoage, T., Ding, Y., Xu, X., 2012. Quantifying cardiac functions in embryonic and adult zebrafish. Methods Mol Biol 843, 11-20. Horb, L.D., Horb, M.E., 2010. BrunoL1 regulates endoderm proliferation through translational enhancement of cyclin A2 mRNA. Dev Biol 345, 156-169. Huichalaf, C.H., Sakai, K., Wang, G.L., Timchenko, N.A., Timchenko, L., 2007. Regulation of the promoter of CUG triplet repeat binding protein, Cugbp1, during myogenesis. Gene 396, 391-402. Huot, M.E., Bisson, N., Davidovic, L., Mazroui, R., Labelle, Y., Moss, T., Khandjian, E.W., 2005. The RNA-binding protein fragile X-related 1 regulates somite formation in Xenopus laevis. Mol Biol Cell 16, 4350-4361. Iakova, P., Wang, G.L., Timchenko, L., Michalak, M., Pereira-Smith, O.M., Smith, J.R., Timchenko, N.A., 2004. Competition of CUGBP1 and calreticulin for the regulation of p21 translation determines cell fate. EMBO J 23, 406-417. James-Zorn, C., Ponferrada, V.G., Jarabek, C.J., Burns, K.A., Segerdell, E.J., Lee, J., Snyder, K., Bhattacharyya, B., Karpinka, J.B., Fortriede, J., Bowes, J.B., Zorn, A.M., Vize, P.D., 2013. Xenbase: expansion and updates of the Xenopus model organism database. Nuc Acids Res 41, D865-870. Jin, Y., Suzuki, H., Maegawa, S., Endo, H., Sugano, S., Hashimoto, K., Yasuda, K., Inoue, K., 2003. A vertebrate RNA-binding protein Fox-1 regulates tissue-specific splicing via the pentanucleotide GCAUG. EMBO J 22, 905-912. Johnston, J.J., Teer, J.K., Cherukuri, P.F., Hansen, N.F., Loftus, S.K., Chong, K., Mullikin, J.C., Biesecker, L.G., 2010. Massively parallel sequencing of exons on the X chromosome identifies RBM10 as the gene that causes a syndromic form of cleft palate. Am J Hum Genet 86, 743-748. Jumaa, H., Wei, G., Nielsen, P.J., 1999. Blastocyst formation is blocked in mouse embryos lacking the splicing factor SRp20. Curr Biol 9, 899-902. Justice, M.J., Hirschi, K.K., 2010. The role of quaking in mammalian embryonic development. Adv Exp Med Biol 693, 82-92. Kalsotra, A., Cooper, T.A., 2011. Functional consequences of developmentally regulated alternative splicing. Nature reviews. Genetics 12, 715-729. Kalsotra, A., Wang, K., Li, P.F., Cooper, T.A., 2010. MicroRNAs coordinate an alternative splicing network during mouse postnatal heart development. Genes Dev. Kalsotra, A., Xiao, X., Ward, A.J., Castle, J.C., Johnson, J.M., Burge, C.B., Cooper, T.A., 2008. A postnatal switch of CELF and MBNL proteins reprograms alternative splicing in the developing heart. Proc Natl Acad Sci U S A 105, 20333-20338. Kanadia, R.N., Johnstone, K.A., Mankodi, A., Lungu, C., Thornton, C.A., Esson, D., Timmers, A.M., Hauswirth, W.W., Swanson, M.S., 2003a. A muscleblind knockout model for myotonic dystrophy. Science 302, 1978-1980. Kanadia, R.N., Shin, J., Yuan, Y., Beattie, S.G., Wheeler, T.M., Thornton, C.A., Swanson, M.S., 2006. Reversal of RNA missplicing and myotonia after muscleblind overexpression 202 in a mouse poly(CUG) model for myotonic dystrophy. Proc Natl Acad Sci U S A 103, 11748-11753. Kanadia, R.N., Urbinati, C.R., Crusselle, V.J., Luo, D., Lee, Y.J., Harrison, J.K., Oh, S.P., Swanson, M.S., 2003b. Developmental expression of mouse muscleblind genes Mbnl1, Mbnl2 and Mbnl3. Gene Expr Patterns 3, 459-462. Kartha, R.V., Subramanian, S., 2014. Competing endogenous RNAs (ceRNAs): new entrants to the intricacies of gene regulation. Front Genet 5, 8. Karunakaran, D.K., Congdon, S., Guerrette, T., Banday, A.R., Lemoine, C., Chhaya, N., Kanadia, R., 2013. The expression analysis of Sfrs10 and Celf4 during mouse retinal development. Gene Expr Patterns 13, 425-436. Kelemen, O., Convertini, P., Zhang, Z., Wen, Y., Shen, M., Falaleeva, M., Stamm, S., 2013. Function of alternative splicing. Gene 514, 1-30. Kent, W.J., 2002. BLAT--the BLAST-like alignment tool. Genome Res 12, 656-664. Keyte, A., Hutson, M.R., 2012. The neural crest in cardiac congenital anomalies. Differentiation 84, 25-40. Kim, E., Magen, A., Ast, G., 2007. Different levels of alternative splicing among eukaryotes. Nuc Acids Res 35, 125-131. Kim, T., Kim, J.O., Oh, J.G., Hong, S.E., Kim do, H., 2014. Pressure-overload cardiac hypertrophy is associated with distinct alternative splicing due to altered expression of splicing factors. Mol Cells 37, 81-87. Kim-Ha, J., Kerr, K., Macdonald, P., 1995. Translational regulation of oskar mRNA by bruno, an ovarian RNA-binding protein, is essential. Cell 81, 403-412. Kirby, M.L., 2007. Cardiac Development. Oxford University Press, New York. Kishore, S., Luber, S., Zavolan, M., 2010. Deciphering the role of RNA-binding proteins in the post-transcriptional control of gene expression. Brief Funct Genomics 9, 391-404. Komamura, K., Iwai, N., Kokame, K., Yasumura, Y., Kim, J., Yamagishi, M., Morisaki, T., Kimura, A., Tomoike, H., Kitakaze, M., Miyatake, K., 2004. The role of a common TNNT2 polymorphism in cardiac hypertrophy. J Hum Genet 49, 129-133. Komiyama, M., Maruyama, K., Shimada, Y., 1990. Assembly of connectin (titin) in relation to myosin and alpha-actinin in cultured cardiac myocytes. J Muscle Res Cell Motil 11, 419-428. Kondo, T., Furuta, T., Mitsunaga, K., Ebersole, T.A., Shichiri, M., Wu, J., Artzt, K., Yamamura, K., Abe, K., 1999. Genomic organization and expression analysis of the mouse qkI locus. Mamm Genome 10, 662-669. Kong, J., Lasko, P., 2012. Translational control in cellular and developmental processes. Nature reviews. Genetics 13, 383-394. Kong, S.W., Hu, Y.W., Ho, J.W., Ikeda, S., Polster, S., John, R., Hall, J.L., Bisping, E., Pieske, B., dos Remedios, C.G., Pu, W.T., 2010. Heart failure-associated changes in RNA splicing of sarcomere genes. Circ Cardiovasc Genet 3, 138-146. Koshelev, M., Sarma, S., Price, R.E., Wehrens, X.H., Cooper, T.A., 2010. Heart-specific overexpression of CUGBP1 reproduces functional and molecular abnormalities of myotonic dystrophy type 1. Hum Mol Genet 19, 1066-1075. Kress, C., Gautier-Courteille, C., Osborne, H.B., Babinet, C., Paillard, L., 2007. Inactivation of CUG-BP1/CELF1 causes growth, viability, and spermatogenesis defects in mice. Mol Cell Biol 27, 1146-1157. 203 Kruithof, B.P., Duim, S.N., Moerkamp, A.T., Goumans, M.J., 2012. TGFbeta and BMP signaling in cardiac cushion formation: Lessons from mice and chicken. Differentiation 84, 89- 102. Kuroyanagi, H., 2009. Fox-1 family of RNA-binding proteins. Cell Mol Life Sci 66, 3895- 3907. Kuyumcu-Martinez, N.M., Wang, G.S., Cooper, T.A., 2007. Increased steady-state levels of CUGBP1 in myotonic dystrophy 1 are due to PKC-mediated hyperphosphorylation. Mol Cell 28, 68-78. Ladd, A., Charlet-B., N., Cooper, T., 2001. The CELF family of RNA binding proteins is implicated in cell-specific and developmentally regulated alternative splicing. Mol Cell Biol 21, 1285-1296. Ladd, A., Cooper, T., 2004. Multiple domains control the subcellular localization and activity of ETR-3, a regulator of nuclear and cytoplasmic RNA processing events. J Cell Sci 117, 3519-3529. Ladd, A., Nguyen, N., Malhotra, K., Cooper, T., 2004. CELF6, a member of the CELF family of RNA binding proteins, regulates muscle-specific splicing enhancer-dependent alternative splicing. J Biol Chem 279, 17756-17764. Ladd, A., Stenberg, M., Swanson, M., Cooper, T., 2005a. A dynamic balance between activation and repression regulates pre-mRNA alternative splicing during heart development. Dev Dyn 233, 783-793. Ladd, A., Taffet, G., Hartley, C., Kearney, D., Cooper, T., 2005b. Cardiac-specific repression of CELF activity disrupts alternative splicing and causes cardiomyopathy. Mol Cell Biol 25, 6267-6278. Ladd, A.N., 2012. CUG-BP, Elav-like family (CELF)-mediated alternative splicing regulation in the brain during health and disease. Mol Cell Neurosci. Ladd, A.N., Yatskievych, T.A., Antin, P.B., 1998. Regulation of avian cardiac myogenesis by activin/TGFbeta and bone morphogenetic proteins. Dev Biol 204, 407-419. Lale, S., Yu, S., Ahmed, A., 2011. Complex congenital heart defects in association with maternal diabetes and partial deletion of the A2BP1 gene. Fetal Pediatr Pathol 30, 161-166. Lasko, P., 2003. Gene regulation at the RNA layer: RNA binding proteins in intercellular signaling networks. Sci STKE 2003, RE6. Lee, J.E., Lee, J.Y., Wilusz, J., Tian, B., Wilusz, C.J., 2010. Systematic analysis of cis-elements in unstable mRNAs demonstrates that CUGBP1 is a key regulator of mRNA decay in muscle cells. PLoS One 5, e11201. Lee, M.H., Schedl, T., 2006. RNA-binding proteins. WormBook, 1-13. Legato, M.J., 1972. Ultrastructural characteristics of the rat ventricular cell grown in tissue culture, with special reference to sarcomerogenesis. J Molec Cell Cardiol 4, 299-317. LeMasters, K.E., Blech-Hermoni, Y., Stillwagon, S.J., Vajda, N.A., Ladd, A.N., 2012. Loss of muscleblind-like 1 promotes invasive mesenchyme formation in endocardial cushions by stimulating autocrine TGFbeta3. BMC Dev Biol 12, 22. Levers, T., Tait, S., Birling, M.-C., Brophy, P., Price, D., 2002. Etr-r3/mNapor, encoding an ELAV-type RNA binding protein, is expressed in differentiating cells in the developing rodent forebrain. Mech Dev 112, 191-193. 204 Li, D., Bachinski, L., Roberts, R., 2001. Genomic organization and isoform-specific tissue expression of human NAPOR (CUGBP2) as a candidate gene for familial arrhythmogenic right ventricular dysplasia. Genomics 74, 396-401. Li, D., Morales, A., Gonzalez-Quintana, J., Norton, N., Siegfried, J.D., Hofmeyer, M., Hershberger, R.E., 2010. Identification of novel mutations in RBM20 in patients with dilated cardiomyopathy. Clin Transl Sci 3, 90-97. Li, S., Guo, W., Dewey, C.N., Greaser, M.L., 2013. Rbm20 regulates titin alternative splicing as a splicing repressor. Nuc Acids Res 41, 2659-2672. Li, X., Wang, J., Manley, J.L., 2005. Loss of splicing factor ASF/SF2 induces G2 cell cycle arrest and apoptosis, but inhibits internucleosomal DNA fragmentation. Genes Dev 19, 2705-2714. Lichtner, P., Attié-Bitach, T., Schuffenhauer, S., Henwood, J., Bouvagnet, P., Scambler, P., Meitinger, T., Vekemans, M., 2002. Expression and mutation analysis of Brunol3, a candidate gene for heart and thymus developmental defects associated with partial monosomy 10p. J Mol Med 80, 431-442. Lin, M.I., Das, I., Schwartz, G.M., Tsoulfas, P., Mikawa, T., Hempstead, B.L., 2000. Trk C receptor signaling regulates cardiac myocyte proliferation during early heart development in vivo. Dev Biol 226, 180-191. Lin, Z.X., Holtzer, S., Schultheiss, T., Murray, J., Masaki, T., Fischman, D.A., Holtzer, H., 1989. Polygons and adhesion plaques and the disassembly and assembly of myofibrils in cardiac myocytes. Journal Cell Biol 108, 2355-2367. Liu, Y., Huang, H., Yuan, B., Luo, T., Li, J., Qin, X., 2014. Suppression of CUGBP1 inhibits growth of hepatocellular carcinoma cells. Clin Invest Med 37, E10-18. Liu, Y.F., Liu, H.Y., Tu, L.C., Lin, C.W., Hsiao, K.M., Pan, H., 2008. Zebrafish muscleblind-like genes: identification, structural features and expression. Comp Biochem Physiol B Biochem Mol Biol 151, 118-124. Liu, Z.P., Nakagawa, O., Nakagawa, M., Yanagisawa, H., Passier, R., Richardson, J.A., Srivastava, D., Olson, E.N., 2001. CHAMP, a novel cardiac-specific helicase regulated by MEF2C. Dev Biol 234, 497-509. Liu, Z.P., Olson, E.N., 2002. Suppression of proliferation and cardiomyocyte hypertrophy by CHAMP, a cardiac-specific RNA helicase. Proc Nat Acad Sci U S A 99, 2043-2048. Lockhart, M., Wirrig, E., Phelps, A., Wessels, A., 2011. Extracellular matrix and heart development. Birth Defects Res A Clin Mol Teratol 91, 535-550. Loria, P., Duke, A., Rand, J., Hobert, O., 2003. Two neuronal, nuclear-localized RNA binding proteins involved in synaptic transmission. Curr Biol 13, 1317-1323. Lovci, M.T., Ghanem, D., Marr, H., Arnold, J., Gee, S., Parra, M., Liang, T.Y., Stark, T.J., Gehman, L.T., Hoon, S., Massirer, K.B., Pratt, G.A., Black, D.L., Gray, J.W., Conboy, J.G., Yeo, G.W., 2013. Rbfox proteins regulate alternative mRNA splicing through evolutionarily conserved RNA bridges. Nat Struct Mol Biol 20, 1434-1442. Lu, J.Y., Schneider, R.J., 2004. Tissue distribution of AU-rich mRNA-binding proteins involved in regulation of mRNA decay. J Biol Chem 279, 12974-12979. Lu, X., Timchenko, N., Timchenko, L., 1999. Cardiac elav-type RNA-binding protein (ETR-3) binds to RNA CUG repeats expanded in myotonic dystrophy. Hum Mol Gen 8, 53-60. Lunde, B.M., Moore, C., Varani, G., 2007. RNA-binding proteins: modular design for efficient function. Nat Rev Mol Cell Biol 8, 479-490. Maas, S., 2010. Gene regulation through RNA editing. Discov Med 10, 379-386. 205 Maas, S., Melcher, T., Herb, A., Seeburg, P.H., Keller, W., Krause, S., Higuchi, M., O'Connell, M.A., 1996. Structural requirements for RNA editing in glutamate receptor pre- mRNAs by recombinant double-stranded RNA adenosine deaminase. J Biol Chem 271, 12221-12226. Machuca-Tzili, L.E., Buxton, S., Thorpe, A., Timson, C.M., Wigmore, P., Luther, P.K., Brook, J.D., 2011. Zebrafish deficient for Muscleblind-like 2 exhibit features of myotonic dystrophy. Dis Models Mech 4, 381-392. MacLellan, W.R., Schneider, M.D., 2000. Genetic dissection of cardiac growth control pathways. Annu Rev Physiol 62, 289-319. Magdaleno, S., Jensen, P., Brumwell, C.L., Seal, A., Lehman, K., Asbury, A., Cheung, T., Cornelius, T., Batten, D.M., Eden, C., Norland, S.M., Rice, D.S., Dosooye, N., Shakya, S., Mehta, P., Curran, T., 2006. BGEM: an in situ hybridization database of gene expression in the embryonic and adult mouse nervous system. PLoS Biol 4, e86. Mallela, A., Nishikura, K., 2012. A-to-I editing of protein coding and noncoding RNAs. Crit Rev Biochem Mol Biol 47, 493-501. Mango, R., Biocca, S., del Vecchio, F., Clementi, F., Sangiuolo, F., Amati, F., Filareto, A., Grelli, S., Spitalieri, P., Filesi, I., Favalli, C., Lauro, R., Mehta, J.L., Romeo, F., Novelli, G., 2005. In vivo and in vitro studies support that a new splicing isoform of OLR1 gene is protective against acute myocardial infarction. Circ Res 97, 152-158. Maniatis, T., Tasic, B., 2002. Alternative pre-mRNA splicing and proteome expansion in metazoans. Nature 418, 236-243. Manner, J., 2000. Cardiac looping in the chick embryo: a morphological review with special reference to terminological and biomechanical aspects of the looping process. Anat Rec 259, 248-262. Manner, J., 2013. On the form problem of embryonic heart loops, its geometrical solutions, and a new biophysical concept of cardiac looping. Ann Anat 195, 312-323. Manner, J., Thrane, L., Norozi, K., Yelbuz, T.M., 2008. High-resolution in vivo imaging of the cross-sectional deformations of contracting embryonic heart loops using optical coherence tomography. Dev Dyn 237, 953-961. Maragh, S., Miller, R.A., Bessling, S.L., McGaughey, D.M., Wessels, M.W., de Graaf, B., Stone, E.A., Bertoli-Avella, A.M., Gearhart, J.D., Fisher, S., McCallion, A.S., 2011. Identification of RNA binding motif proteins essential for cardiovascular development. BMC Dev Biol 11, 62. Marquis, J., Paillard, L., Audic, Y., Cosson, B., Danos, O., Le Bec, C., Osborne, H.B., 2006. CUG- BP1/CELF1 requires UGU-rich sequences for high-affinity binding. Biochem J 400, 291-301. Masuda, A., Andersen, H.S., Doktor, T.K., Okamoto, T., Ito, M., Andresen, B.S., Ohno, K., 2012. CUGBP1 and MBNL1 preferentially bind to 3' UTRs and facilitate mRNA decay. Sci Rep 2, 209. Masuda, K., Marasa, B., Martindale, J.L., Halushka, M.K., Gorospe, M., 2009. Tissue- and age- dependent expression of RNA-binding proteins that influence mRNA turnover and translation. Aging (Albany NY) 1, 681-698. Matsui, T., Sasaki, A., Akazawa, N., Otani, H., Bessho, Y., 2012. Celf1 regulation of dmrt2a is required for somite symmetry and left-right patterning during zebrafish development. Development 139, 3553-3560. 206 McAuliffe, J., Gao, L., Solaro, R., 1990. Changes in myofibrillar activation and troponin C Ca2+ binding associated with troponin T isoform switching in developing rabbit heart. Circ Res 66, 1204-1216. McKee, A.E., Minet, E., Stern, C., Riahi, S., Stiles, C.D., Silver, P.A., 2005. A genome-wide in situ hybridization map of RNA-binding proteins reveals anatomically restricted expression in the developing mouse brain. BMC Dev Biol 5, 14. Medioni, C., Astier, M., Zmojdzian, M., Jagla, K., Semeriva, M., 2008. Genetic control of cell morphogenesis during Drosophila melanogaster cardiac tube formation. Journal Cell Biol 182, 249-261. Medioni, C., Mowry, K., Besse, F., 2012. Principles and roles of mRNA localization in animal development. Development 139, 3263-3276. Meins, M., Schlickum, S., Wilhelm, C., Missbach, J., Yadav, S., Glaser, B., Grzmil, M., Burfeind, P., Laccone, F., 2002. Identification and characterization of murine Brunol4, a new member of the elav/bruno family. Cytogenet Genome Res 97, 254-260. Michalowski, S., Miller, J., Urbinati, C., Paliouras, M., Swanson, M., Griffith, J., 1999. Visualization of double-stranded RNAs from the myotonic dystrophy protein kinase gene and interactions with CUG-binding protein. Nuc Acids Res 27, 3534-3542. Mientjes, E.J., Willemsen, R., Kirkpatrick, L.L., Nieuwenhuizen, I.M., Hoogeveen-Westerveld, M., Verweij, M., Reis, S., Bardoni, B., Hoogeveen, A.T., Oostra, B.A., Nelson, D.L., 2004. Fxr1 knockout mice show a striated muscle phenotype: implications for Fxr1p function in vivo. Hum Mol Genet 13, 1291-1302. Miller, R.A., Christoforou, N., Pevsner, J., McCallion, A.S., Gearhart, J.D., 2008. Efficient array- based identification of novel cardiac genes through differentiation of mouse ESCs. PLoS One 3, e2176. Milne, C., Hodgkin, J., 1999. ETR-1, a homologue of a protein linked to myotonic dystropy, is essential for muscle development in Caenorhabditis elegans. Curr Biol 9, 1243-1246. Misquitta, C.M., Chen, T., Grover, A.K., 2006. Control of protein expression through mRNA stability in calcium signalling. Cell Calcium 40, 329-346. Misquitta, C.M., Iyer, V.R., Werstiuk, E.S., Grover, A.K., 2001. The role of 3'-untranslated region (3'-UTR) mediated mRNA stability in cardiovascular pathophysiology. Mol Cell Biochem 224, 53-67. Modrek, B., Lee, C., 2002. A genomic view of alternative splicing. Nat Genet 30, 13-19. Moorman, A.F., Christoffels, V.M., 2003. Cardiac chamber formation: development, genes, and evolution. Physiol Rev 83, 1223-1267. Moraes, K.C., Monteiro, C.J., Pacheco-Soares, C., 2013. A novel function for CUGBP2 in controlling the pro-inflammatory stimulus in H9c2 cells: subcellular trafficking of messenger molecules. Cell Biol Int 37, 1129-1138. Moraes, K.C., Wilusz, C.J., Wilusz, J., 2006. CUG-BP binds to RNA substrates and recruits PARN deadenylase. RNA 12, 1084-1091. Mori, A.D., Bruneau, B.G., 2004. TBX5 mutations and congenital heart disease: Holt-Oram syndrome revealed. Curr Opin Cardiol 19, 211-215. Mori, D., Sasagawa, N., Kino, Y., Ishiura, S., 2008a. Quantitative analysis of CUG-BP1 binding to RNA repeats. J Biochem 143, 377-383. Mueller, C.F., Berger, A., Zimmer, S., Tiyerili, V., Nickenig, G., 2009. The heterogenous nuclear riboprotein S1-1 regulates AT1 receptor gene expression via transcriptional and posttranscriptional mechanisms. Arch Biochem Biophys 488, 76-82. 207 Mukhopadhyay, D., Houchen, C., Kennedy, S., Dieckgraefe, B., Anant, S., 2003a. Coupled mRNA stabilization and translational silencing of cyclooxygenase-2 by a novel RNA binding protein, CUGBP2. Mol Cell 11, 113-126. Mukhopadhyay, D., Jung, J., Murmu, N., Houchen, C.W., Dieckgraefe, B.K., Anant, S., 2003b. CUGBP2 plays a critical role in apoptosis of breast cancer cells in response to genotoxic injury. Ann N Y Acad Sci 1010, 504-509. Murmu, N., Jung, J., Mukhopadhyay, D., Houchen, C.W., Riehl, T.E., Stenson, W.F., Morrison, A.R., Arumugam, T., Dieckgraefe, B.K., Anant, S., 2004. Dynamic antagonism between RNA-binding protein CUGBP2 and cyclooxygenase-2-mediated prostaglandin E2 in radiation damage. Proc Natl Acad Sci U S A 101, 13873-13878. Naha, N., Lee, H.Y., Naser, M.I., Park, T.J., Kim, S.H., Kim, M.O., 2009. Ethanol inhibited apoptosis-related RNA binding protein, Napor-3 gene expression in the prenatal rat brain. Med Sci Monit 15, BR6-12. Natarajan, G., Ramalingam, S., Ramachandran, I., May, R., Queimado, L., Houchen, C.W., Anant, S., 2008. CUGBP2 Downregulation By Prostaglandin E2 Protects Colon Cancer Cells from Radiation-Induced Mitotic Catastrophe. Am J Physiol Gastrointest Liver Physiol. Naya, F.J., Black, B.L., Wu, H., Bassel-Duby, R., Richardson, J.A., Hill, J.A., Olson, E.N., 2002. Mitochondrial deficiency and cardiac sudden death in mice lacking the MEF2A transcription factor. Nat Med 8, 1303-1309. Nelson, T.J., Balza, R., Jr., Xiao, Q., Misra, R.P., 2005. SRF-dependent gene expression in isolated cardiomyocytes: regulation of genes involved in cardiac hypertrophy. J Molec Cell Cardiol 39, 479-489. Nilsen, T.W., Graveley, B.R., 2010. Expansion of the eukaryotic proteome by alternative splicing. Nature 463, 457-463. Nir, R., Grossman, R., Paroush, Z., Volk, T., 2012. Phosphorylation of the Drosophila melanogaster RNA-binding protein HOW by MAPK/ERK enhances its dimerization and activity. PLoS Genet 8, e1002632. Ocorr, K., Fink, M., Cammarato, A., Bernstein, S., Bodmer, R., 2009. Semi-automated Optical Heartbeat Analysis of small hearts. J Vis Exp. Okano, H.J., Darnell, R.B., 1997. A hierarchy of Hu RNA binding proteins in developing and adult neurons. J Neurosci 17, 3024-3037. Ono, K., Kuwabara, Y., Han, J., 2011. MicroRNAs and cardiovascular diseases. FEBS J 278, 1619-1633. Otsuka, N., Tsuritani, K., Sakurai, T., Kato, K., Matoba, R., Itoh, J., Okuyama, S., Yamada, K., Yoneda, Y., 2009. Transcriptional induction and translational inhibition of Arc and Cugbp2 in mice hippocampus after transient global ischemia under normothermic condition. Brain Res 1287, 136-145. Pacini, A., Toscano, A., Cesati, V., Cozzi, A., Meli, E., Di Cesare Mannelli, L., Paternostro, F., Pacini, P., Pellegrini-Giampietro, D.E., 2005. NAPOR-3 RNA binding protein is required for apoptosis in hippocampus. Brain Res Mol Brain Res 140, 34-44. Paillard, L., Legagneux, V., Maniey, D., Osborne, H., 2002. c-Jun ARE targets mRNA deadenlyation by an EDEN-BP (embryo deadenylation element-binding protein)- dependent pathway. J Biol Chem 277, 3232-3235. Paillard, L., Legagneux, V., Osborne, H., 2003. A functional deadenylation assay identifies human CUG-BP as a deadenylation factor. Biol Cell 95, 107-113. 208 Paillard, L., Omilli, F., Legagneux, V., Bassez, T., Maniey, D., Osborne, H., 1998. EDEN and EDEN-BP, a cis element and an associated factor that mediate sequence-specific mRNA deadenylation in Xenopus embryos. EMBO J 17, 278-287. Paillard, L., Osborne, H.B., 2003. East of EDEN was a poly(A) tail. Biol Cell 95, 211-219. Palacios, I.M., 2013. Nonsense-mediated mRNA decay: from mechanistic insights to impacts on human health. Brief Funct Genomics 12, 25-36. Palmer, A.R., 2004. Symmetry breaking and the evolution of development. Science 306, 828-833. Pan, Q., Shai, O., Lee, L.J., Frey, B.J., Blencowe, B.J., 2008. Deep surveying of alternative splicing complexity in the human transcriptome by high-throughput sequencing. Nat Genet 40, 1413-1415. Paris, J., Philippe, M., 1990. Poly(A) metabolism and polysomal recruitment of maternal mRNAs during early Xenopus development. Dev Biol 140, 221-224. Park, J.Y., Li, W., Zheng, D., Zhai, P., Zhao, Y., Matsuda, T., Vatner, S.F., Sadoshima, J., Tian, B., 2011. Comparative analysis of mRNA isoform expression in cardiac hypertrophy and development reveals multiple post-transcriptional regulatory modules. PLoS One 6, e22391. Parra, M.K., Gallagher, T.L., Amacher, S.L., Mohandas, N., Conboy, J.G., 2012. Deep intron elements mediate nested splicing events at consecutive AG dinucleotides to regulate alternative 3' splice site choice in vertebrate 4.1 genes. Mol Cell Biol 32, 2044-2053. Pascual, M., Vicente, M., Monferrer, L., Artero, R., 2006. The Muscleblind family of proteins: an emerging class of regulators of developmentally programmed alternative splicing. Differentiation 74, 65-80. Pende, A., Tremmel, K.D., DeMaria, C.T., Blaxall, B.C., Minobe, W.A., Sherman, J.A., Bisognano, J.D., Bristow, M.R., Brewer, G., Port, J., 1996. Regulation of the mRNA-binding protein AUF1 by activation of the beta-adrenergic receptor signal transduction pathway. J Biol Chem 271, 8493-8501. Perrone-Bizzozero, N., Bolognani, F., 2002. Role of HuD and other RNA-binding proteins in neural development and plasticity. J Neurosci Res 68, 121-126. Person, A.D., Klewer, S.E., Runyan, R.B., 2005. Cell biology of cardiac cushion development. Int Rev Cytol 243, 287-335. Philippar, U., Schratt, G., Dieterich, C., Muller, J.M., Galgoczy, P., Engel, F.B., Keating, M.T., Gertler, F., Schule, R., Vingron, M., Nordheim, A., 2004. The SRF target gene Fhl2 antagonizes RhoA/MAL-dependent activation of SRF. Molecular cell 16, 867-880. Philips, A., Timchenko, L., Cooper, T., 1998. Disruption of splicing regulated by a CUG- binding protein in myotonic dystrophy. Science 280, 737-741. Pickrell, J.K., Pai, A.A., Gilad, Y., Pritchard, J.K., 2010. Noisy splicing drives mRNA isoform diversity in human cells. PLoS Genet 6, e1001236. Poon, K.L., Tan, K.T., Wei, Y.Y., Ng, C.P., Colman, A., Korzh, V., Xu, X.Q., 2012. RNA-binding protein RBM24 is required for sarcomere assembly and heart contractility. Cardiovasc Res 94, 418-427. Pratt, C.A., Mowry, K.L., 2013. Taking a cellular road-trip: mRNA transport and anchoring. Curr Opin Cell Biol 25, 99-106. Pritsker, M., Doniger, T.T., Kramer, L.C., Westcot, S.E., Lemischka, I.R., 2005. Diversification of stem cell molecular repertoire by alternative splicing. Proc Nat Acad Sci U S A 102, 14290-14295. 209 Qayyum, S.R., Webb, S., Anderson, R.H., Verbeek, F.J., Brown, N.A., Richardson, M.K., 2001. Septation and valvar formation in the outflow tract of the embryonic chick heart. Anat Rec 264, 273-283. Raffel, G.D., Chu, G.C., Jesneck, J.L., Cullen, D.E., Bronson, R.T., Bernard, O.A., Gilliland, D.G., 2009. Ott1 (Rbm15) is essential for placental vascular branching morphogenesis and embryonic development of the heart and spleen. Mol Cell Biol 29, 333-341. Raghavan, A., Ogilvie, R.L., Reilly, C., Abelson, M.L., Raghavan, S., Vasdewani, J., Krathwohl, M., Bohjanen, P.R., 2002. Genome-wide analysis of mRNA decay in resting and activated primary human T lymphocytes. Nuc Acids Res 30, 5529-5538. Ramalingam, S., Natarajan, G., Schafer, C., Subramaniam, D., May, R., Ramachandran, I., Queimado, L., Houchen, C.W., Anant, S., 2008. Novel intestinal splice variants of RNA- binding protein CUGBP2: isoform-specific effects on mitotic catastrophe. Am J Physiol Gastrointest Liver Physiol 294, G971-981. Rappsilber, J., Ryder, U., Lamond, A.I., Mann, M., 2002. Large-scale proteomic analysis of the human spliceosome. Genome Res 12, 1231-1245. Rattenbacher, B., Beisang, D., Wiesner, D.L., Jeschke, J.C., von Hohenberg, M., St Louis- Vlasova, I.A., Bohjanen, P.R., 2010. Analysis of CUGBP1 targets identifies GU-repeat sequences that mediate rapid mRNA decay. Mol Cell Biol 30, 3970-3980. Refaat, M.M., Lubitz, S.A., Makino, S., Islam, Z., Frangiskakis, J.M., Mehdi, H., Gutmann, R., Zhang, M.L., Bloom, H.L., MacRae, C.A., Dudley, S.C., Shalaby, A.A., Weiss, R., McNamara, D.M., London, B., Ellinor, P.T., 2012. Genetic variation in the alternative splicing regulator RBM20 is associated with dilated cardiomyopathy. Heart Rhythm 9, 390-396. Reinke, L.M., Xu, Y., Cheng, C., 2012. Snail represses the splicing regulator epithelial splicing regulatory protein 1 to promote epithelial-mesenchymal transition. J Biol Chem 287, 36435-36442. Revil, T., Jerome-Majewska, L.A., 2013. During embryogenesis, esrp1 expression is restricted to a subset of epithelial cells and is associated with splicing of a number of developmentally important genes. Dev Dyn 242, 281-290. Richard, S., Torabi, N., Franco, G.V., Tremblay, G.A., Chen, T., Vogel, G., Morel, M., Cleroux, P., Forget-Richard, A., Komarova, S., Tremblay, M.L., Li, W., Li, A., Gao, Y.J., Henderson, J.E., 2005. Ablation of the Sam68 RNA binding protein protects mice from age- related bone loss. PLoS Genet 1, e74. Roberts, R., Timchenko, N., Miller, J., Reddy, S., Caskey, C., Swanson, M., Timchenko, L., 1997. Altered phosphorylation and intracellular distribution of a (CUG)n triplet repeat RNA-binding protein in patients with myotonic dystrophy and in myotonin protein kinase knockout mice. Proc Natl Acad Sci U S A 94, 13221-13226. Rosenthal, J.J., Seeburg, P.H., 2012. A-to-I RNA editing: effects on proteins key to neural excitability. Neuron 74, 432-439. Rozario, T., DeSimone, D.W., 2010. The extracellular matrix in development and morphogenesis: a dynamic view. Deve Biol 341, 126-140. Rudy, D.E., Yatskievych, T.A., Antin, P.B., Gregorio, C.C., 2001. Assembly of thick, thin, and titin filaments in chick precardiac explants. Dev Dyn 221, 61-71. Rueter, S.M., Dawson, T.R., Emeson, R.B., 1999. Regulation of alternative splicing by RNA editing. Nature 399, 75-80. 210 Ryan, K., Cooper, T., 1996. Muscle-specific splicing enhancers regulate inclusion of the cardiac troponin T alternative exon in embryonic skeletal muscle. Mol Cell Biol 16, 4014-4023. Ryder, S.P., Williamson, J.R., 2004. Specificity of the STAR/GSG domain protein Qk1: implications for the regulation of myelination. RNA 10, 1449-1458. Saba, Z., Nassar, R., Ungerleider, R.M., Oakeley, A.E., Anderson, P.A., 1996. Cardiac troponin T isoform expression correlates with pathophysiological descriptors in patients who underwent corrective surgery for congenital heart disease. Circulation 94, 472-476. Salisbury, E., Sakai, K., Schoser, B., Huichalaf, C., Schneider-Gold, C., Nguyen, H., Wang, G.L., Albrecht, J.H., Timchenko, L.T., 2008. Ectopic expression of cyclin D3 corrects differentiation of DM1 myoblasts through activation of RNA CUG-binding protein, CUGBP1. Exp Cell Res 314, 2266-2278. Salomonis, N., Nelson, B., Vranizan, K., Pico, A.R., Hanspers, K., Kuchinsky, A., Ta, L., Mercola, M., Conklin, B.R., 2009. Alternative splicing in the differentiation of human embryonic stem cells into cardiac precursors. PLoS Comput Biol 5, e1000553. Salomonis, N., Schlieve, C.R., Pereira, L., Wahlquist, C., Colas, A., Zambon, A.C., Vranizan, K., Spindler, M.J., Pico, A.R., Cline, M.S., Clark, T.A., Williams, A., Blume, J.E., Samal, E., Mercola, M., Merrill, B.J., Conklin, B.R., 2010. Alternative splicing regulates mouse embryonic stem cell pluripotency and differentiation. Proc Nat Acad Sci U S A 107, 10514-10519. Sanger, J.W., Wang, J., Fan, Y., White, J., Sanger, J.M., 2010. Assembly and dynamics of myofibrils. J Biomed Biotechnol 2010, 858606. Savkur, R., Phillips, A., Cooper, T., 2001. Aberrant regulation of insulin receptor alternative splicing is associated with insulin resistance in myotonic dystrophy. Nat Genet 29, 40-47. Schneider, C.A., Rasband, W.S., Eliceiri, K.W., 2012. NIH Image to ImageJ: 25 years of image analysis. Nat Methods 9, 671-675. Schoenberg, D.R., Maquat, L.E., 2012. Regulation of cytoplasmic mRNA decay. Nature reviews. Genetics 13, 246-259. Schoser, B., Timchenko, L., 2010. Myotonic dystrophies 1 and 2: complex diseases with complex mechanisms. Curr Genomics 11, 77-90. Seitz, H., Zamore, P.D., 2006. Rethinking the microprocessor. Cell 125, 827-829. Shapiro, I.M., Cheng, A.W., Flytzanis, N.C., Balsamo, M., Condeelis, J.S., Oktay, M.H., Burge, C.B., Gertler, F.B., 2011. An EMT-driven alternative splicing program occurs in human breast cancer and modulates cellular phenotype. PLoS Genet 7, e1002218. Shen, X., Franzini-Armstrong, C., Lopez, J.R., Jones, L.R., Kobayashi, Y.M., Wang, Y., Kerrick, W.G., Caswell, A.H., Potter, J.D., Miller, T., Allen, P.D., Perez, C.F., 2007. Triadins modulate intracellular Ca(2+) homeostasis but are not essential for excitation- contraction coupling in skeletal muscle. J Biol Chem 282, 37864-37874. Shepard, P.J., Hertel, K.J., 2009. The SR protein family. Genome Biol 10, 242. Shibata, H., Huynh, D.P., Pulst, S.M., 2000. A novel protein with RNA-binding motifs interacts with ataxin-2. Hum Molec Genet 9, 1303-1313. Shin, C., Feng, Y., Manley, J.L., 2004. Dephosphorylated SRp38 acts as a splicing repressor in response to heat shock. Nature 427, 553-558. Siedner, S., Kruger, M., Schroeter, M., Metzler, D., Roell, W., Fleischmann, B.K., Hescheler, J., Pfitzer, G., Stehle, R., 2003. Developmental changes in contractility and sarcomeric 211 proteins from the early embryonic to the adult stage in the mouse heart. J Physiol 548, 493-505. Singh, G., Charlet, B.N., Han, J., Cooper, T.A., 2004. ETR-3 and CELF4 protein domains required for RNA binding and splicing activity in vivo. Nuc Acids Res 32, 1232-1241. Siomi, H., Dreyfuss, G., 1997. RNA-binding proteins as regulators of gene expression. Curr Opin Genet Dev 7, 345-353. Siomi, M.C., Siomi, H., Sauer, W.H., Srinivasan, S., Nussbaum, R.L., Dreyfuss, G., 1995. FXR1, an autosomal homolog of the fragile X mental retardation gene. EMBO J 14, 2401- 2408. Song, H.K., Hong, S.E., Kim, T., Kim do, H., 2012. Deep RNA sequencing reveals novel cardiac transcriptomic signatures for physiological and pathological hypertrophy. PLoS One 7, e35552. St Pierre, S.E., Ponting, L., Stefancsik, R., McQuilton, P., 2014. FlyBase 102--advanced approaches to interrogating FlyBase. Nuc Acids Res 42, D780-788. Sugnet, C.W., Srinivasan, K., Clark, T.A., O'Brien, G., Cline, M.S., Wang, H., Williams, A., Kulp, D., Blume, J.E., Haussler, D., Ares, M., Jr., 2006. Unusual intron conservation near tissue-regulated exons found by splicing microarrays. PLoS Comput Biol 2, e4. Sun, W., Wagnon, J.L., Mahaffey, C.L., Briese, M., Ule, J., Frankel, W.N., 2013. Aberrant sodium channel activity in the complex seizure disorder of Celf4 mutant mice. J Physiol 591, 241-255. Sureban, S.M., Murmu, N., Rodriguez, P., May, R., Maheshwari, R., Dieckgraefe, B.K., Houchen, C.W., Anant, S., 2007. Functional antagonism between RNA binding proteins HuR and CUGBP2 determines the fate of COX-2 mRNA translation. Gastroenterology 132, 1055-1065. Sutherland, L.C., Rintala-Maki, N.D., White, R.D., Morin, C.D., 2005. RNA binding motif (RBM) proteins: a novel family of apoptosis modulators? J Cell Biochem 94, 5-24. Suzuki, H., Jin, Y., Otani, H., Yasuda, K., Inoue, K., 2002. Regulation of alternative splicing of α-actinin transcript by Bruno-like proteins. Genes Cells 7, 133-141. Suzuki, H., Maegawa, S., Nishibu, T., Sugiyama, T., Yasuda, K., Inoue, K., 2000. Vegetal localization of the maternal mRNA encoding an EDEN-BP/Bruno-like protein in zebrafish. Mech Dev 93, 205-209. Suzuki, H., Takeuchi, M., Sugiyama, A., Alam, A.K., Vu, L.T., Sekiyama, Y., Dam, H.C., Ohki, S.Y., Tsukahara, T., 2012. Alternative splicing produces structural and functional changes in CUGBP2. BMC Biochem 13, 6. Tahara, N., Bessho, Y., Matsui, T., 2013. Celf1 is required for formation of endoderm-derived organs in zebrafish. Int J Mol Sci 14, 18009-18023. Takahashi, N., Sasagawa, N., Suzuki, K., Ishiura, S., 2000. The CUG-binding protein binds specifically to UG dinucleotide repeats in a yeast three-hybird system. Biochem Biophys Res Comm 277, 518-523. Takahashi, N., Sasagawa, N., Usuki, F., Kino, Y., Kawahara, H., Sorimachi, H., Maeda, T., Suzuki, K., Ishiura, S., 2001. Coexpression of the CUG-binding protein reduces DM protein kinase expression in COS cells. J Biochem 130, 581-587. Talwar, S., Balasubramanian, S., Sundaramurthy, S., House, R., Wilusz, C.J., Kuppuswamy, D., D'Silva, N., Gillespie, M.B., Hill, E.G., Palanisamy, V., 2013. Overexpression of RNA- binding protein CELF1 prevents apoptosis and destabilizes pro-apoptotic mRNAs in oral cancer cells. RNA Biol 10, 277-286. 212 Teplova, M., Song, J., Gaw, H.Y., Teplov, A., Patel, D.J., 2010. Structural insights into RNA recognition by the alternate-splicing regulator CUG-binding protein 1. Structure 18, 1364-1377. Terai, M., Komiyama, M., Shimada, Y., 1989. Myofibril assembly is linked with vinculin, alpha-actinin, and cell-substrate contacts in embryonic cardiac myocytes in vitro. Cell Motil Cytoskeleton 12, 185-194. Terenzi, F., Brimacombe, K.R., Penn, M.S., Ladd, A.N., 2009. CELF-mediated alternative splicing is required for cardiac function during early, but not later, postnatal life. J Mol Cell Cardiol 46, 395-404. Terenzi, F., Ladd, A.N., 2010. Conserved developmental alternative splicing of muscleblind- like (MBNL) transcripts regulates MBNL localization and activity. RNA Biol 7, 43-55. Timchenko, L., Miller, J., Timchenko, N., Devore, D., Datar, K., Lin, L., Roberts, R., Caskey, C., Swanson, M., 1996a. Identification of a (CUG)n triplet repeat RNA-binding protein and its expression in myotonic dystrophy. Nuc Acids Res 24, 4407-4414. Timchenko, L., Timchenko, N., Caskey, C., Roberts, R., 1996b. Novel proteins with binding specificity for DNA CTG repeats and RNA CUG repeats - implications for myotonic dystrophy. Hum Mol Gen 5, 115-121. Timchenko, L.T., Salisbury, E., Wang, G.L., Nguyen, H., Albrecht, J.H., Hershey, J.W., Timchenko, N.A., 2006. Age-specific CUGBP1-eIF2 complex increases translation of CCAAT/enhancer-binding protein beta in old liver. J Biol Chem 281, 32806-32819. Timchenko, N., Cai, Z.-J., Welm, A., Reddy, S., Ashizawa, T., Timchenko, L., 2001a. RNA CUG repeats sequester CUGBP1 and alter protein levels and activity of CUGBP1. J Biol Chem 276, 7820-7826. Timchenko, N., Iakova, P., Cai, Z.-J., Smith, J., Timchenko, L., 2001b. Molecular basis for impaired muscle differentiation in myotonic dystrophy. Mol Cell Biol 21, 6927-6938. Timchenko, N.A., Patel, R., Iakova, P., Cai, Z.J., Quan, L., Timchenko, L.T., 2004. Overexpression of CUG triplet repeat-binding protein, CUGBP1, in mice inhibits myogenesis. J Biol Chem 279, 13129-13139. Timchenko, N.A., Wang, G.L., Timchenko, L.T., 2005. RNA CUG-binding protein 1 increases translation of 20-kDa isoform of CCAAT/enhancer-binding protein beta by interacting with the alpha and beta subunits of eukaryotic initiation translation factor 2. J Biol Chem 280, 20549-20557. Timchenko, N.A., Welm, A.L., Lu, X., Timchenko, L.T., 1999. CUG repeat binding protein (CUGBP1) interacts with the 5' region of C/EBPbeta mRNA and regulates translation of C/EBPbeta isoforms. Nuc Acids Res 27, 4517-4525. Tsuda, K., Kuwasako, K., Takahashi, M., Someya, T., Inoue, M., Terada, T., Kobayashi, N., Shirouzu, M., Kigawa, T., Tanaka, A., Sugano, S., Guntert, P., Muto, Y., Yokoyama, S., 2009. Structural basis for the sequence-specific RNA-recognition mechanism of human CUG-BP1 RRM3. Nuc Acids Res 37, 5151-5166. Twyffels, L., Gueydan, C., Kruys, V., 2011. Shuttling SR proteins: more than splicing factors. FEBS J 278, 3246-3255. Ueyama, T., Kasahara, H., Ishiwata, T., Yamasaki, N., Izumo, S., 2003. Csm, a cardiac-specific isoform of the RNA helicase Mov10l1, is regulated by Nkx2.5 in embryonic heart. J Biol Chem 278, 28750-28757. Ule, J., Jensen, K., Mele, A., Darnell, R.B., 2005. CLIP: a method for identifying protein-RNA interaction sites in living cells. Methods 37, 376-386. 213 Ule, J., Jensen, K.B., Ruggiu, M., Mele, A., Ule, A., Darnell, R.B., 2003. CLIP identifies Nova- regulated RNA networks in the brain. Science 302, 1212-1215. Vajda, N.A., Brimacombe, K.R., LeMasters, K.E., Ladd, A.N., 2009. Muscleblind-like 1 is a negative regulator of TGF-beta-dependent epithelial-mesenchymal transition of atrioventricular canal endocardial cells. Dev Dyn 238, 3266-3272. Van't Padje, S., Chaudhry, B., Severijnen, L.A., van der Linde, H.C., Mientjes, E.J., Oostra, B.A., Willemsen, R., 2009. Reduction in fragile X related 1 protein causes cardiomyopathy and muscular dystrophy in zebrafish. J Exp Biol 212, 2564-2570. Venables, J.P., Vignal, E., Baghdiguian, S., Fort, P., Tazi, J., 2012. Tissue-specific alternative splicing of Tak1 is conserved in deuterostomes. Mol Biol Evol 29, 261-269. Verma, S.K., Deshmukh, V., Liu, P., Nutter, C.A., Espejo, R., Hung, M.L., Wang, G.S., Yeo, G.W., Kuyumcu-Martinez, M.N., 2013. Reactivation of fetal splicing programs in diabetic hearts is mediated by protein kinase C signaling. J Biol Chem 288, 35372-35386. Vitelli, F., Morishima, M., Taddei, I., Lindsay, E.A., Baldini, A., 2002. Tbx1 mutation causes multiple cardiovascular defects and disrupts neural crest and cranial nerve migratory pathways. Hum Mol Genet 11, 915-922. Vlasova, I.A., Bohjanen, P.R., 2008. Posttranscriptional regulation of gene networks by GU- rich elements and CELF proteins. RNA Biol 5, 201-207. Vlasova, I.A., Tahoe, N.M., Fan, D., Larsson, O., Rattenbacher, B., Sternjohn, J.R., Vasdewani, J., Karypis, G., Reilly, C.S., Bitterman, P.B., Bohjanen, P.R., 2008. Conserved GU-rich elements mediate mRNA decay by binding to CUG-binding protein 1. Mol Cell 29, 263-270. Vlasova-St Louis, I., Bohjanen, P.R., 2011. Coordinate regulation of mRNA decay networks by GU-rich elements and CELF1. Curr Opin Genet Dev 21, 444-451. Wagner, M., Siddiqui, M.A., 2007. Signal transduction in early heart development (II): ventricular chamber specification, trabeculation, and heart valve formation. Exp Biol Med (Maywood) 232, 866-880. Wagnon, J.L., Briese, M., Sun, W., Mahaffey, C.L., Curk, T., Rot, G., Ule, J., Frankel, W.N., 2012. CELF4 regulates translation and local abundance of a vast set of mRNAs, including genes associated with regulation of synaptic function. PLoS Genet 8, e1003067. Wagnon, J.L., Mahaffey, C.L., Sun, W., Yang, Y., Chao, H.T., Frankel, W.N., 2011. Etiology of a genetically complex seizure disorder in Celf4 mutant mice. Genes Brain Behav 10, 765-777. Wang, E.T., Cody, N.A., Jog, S., Biancolella, M., Wang, T.T., Treacy, D.J., Luo, S., Schroth, G.P., Housman, D.E., Reddy, S., Lecuyer, E., Burge, C.B., 2012. Transcriptome-wide Regulation of Pre-mRNA Splicing and mRNA Localization by Muscleblind Proteins. Cell 150, 710-724. Wang, E.T., Sandberg, R., Luo, S., Khrebtukova, I., Zhang, L., Mayr, C., Kingsmore, S.F., Schroth, G.P., Burge, C.B., 2008. Alternative isoform regulation in human tissue transcriptomes. Nature 456, 470-476. Wang, G.S., Kearney, D.L., De Biasi, M., Taffet, G., Cooper, T.A., 2007. Elevation of RNA- binding protein CUGBP1 is an early event in an inducible heart-specific mouse model of myotonic dystrophy. J Clin Invest 117, 2802-2811. Wang, H.Y., Xu, X., Ding, J.H., Bermingham, J.R., Jr., Fu, X.D., 2001. SC35 plays a role in T cell development and alternative splicing of CD45. Mol Cell 7, 331-342. 214 Wang, J., Takagaki, Y., Manley, J.L., 1996. Targeted disruption of an essential vertebrate gene: ASF/SF2 is required for cell viability. Genes Dev 10, 2588-2599. Wang, Z., Gerstein, M., Snyder, M., 2009. RNA-Seq: a revolutionary tool for transcriptomics. Nat Rev Genet 10, 57-63. Ward, A.J., Cooper, T.A., 2010. The pathobiology of splicing. J Pathol 220, 152-163. Ward, A.J., Rimer, M., Killian, J.M., Dowling, J.J., Cooper, T.A., 2010. CUGBP1 overexpression in mouse skeletal muscle reproduces features of myotonic dystrophy type 1. Hum Mol Genet 19, 3614-3622. Warf, M.B., Nakamori, M., Matthys, C.M., Thornton, C.A., Berglund, J.A., 2009. Pentamidine reverses the splicing defects associated with myotonic dystrophy. Proc Nat Acad Sci U S A 106, 18551-18556. Warkman, A.S., Whitman, S.A., Miller, M.K., Garriock, R.J., Schwach, C.M., Gregorio, C.C., Krieg, P.A., 2012. Developmental expression and cardiac transcriptional regulation of Myh7b, a third myosin heavy chain in the vertebrate heart. Cytoskeleton (Hoboken) 69, 324-335. Warzecha, C.C., Jiang, P., Amirikian, K., Dittmar, K.A., Lu, H., Shen, S., Guo, W., Xing, Y., Carstens, R.P., 2010. An ESRP-regulated splicing programme is abrogated during the epithelial-mesenchymal transition. EMBO J 29, 3286-3300. Warzecha, C.C., Sato, T.K., Nabet, B., Hogenesch, J.B., Carstens, R.P., 2009a. ESRP1 and ESRP2 are epithelial cell-type-specific regulators of FGFR2 splicing. Mol Cell 33, 591-601. Warzecha, C.C., Shen, S., Xing, Y., Carstens, R.P., 2009b. The epithelial splicing factors ESRP1 and ESRP2 positively and negatively regulate diverse types of alternative splicing events. RNA Biol 6, 546-562. Weis, B.L., Schleiff, E., Zerges, W., 2013. Protein targeting to subcellular organelles via MRNA localization. Biochim Biophys Acta 1833, 260-273. Wessels A, M.R., 2000. Cardiac Morphogenesis and Dysmorphogenesis. Humana Press Inc., Totowa, NJ. Wheeler, T.M., Sobczak, K., Lueck, J.D., Osborne, R.J., Lin, X., Dirksen, R.T., Thornton, C.A., 2009. Reversal of RNA dominance by displacement of protein sequestered on triplet repeat RNA. Science 325, 336-339. White, E.J., Brewer, G., Wilson, G.M., 2013. Post-transcriptional control of gene expression by AUF1: mechanisms, physiological targets, and regulation. Biochim Biophys Acta 1829, 680-688. Whitman, S.A., Cover, C., Yu, L., Nelson, D.L., Zarnescu, D.C., Gregorio, C.C., 2011. Desmoplakin and talin2 are novel mRNA targets of fragile X-related protein-1 in cardiac muscle. Circ Res 109, 262-271. Willems, E., Leyns, L., Vandesompele, J., 2008. Standardization of real-time PCR gene expression data from independent biological replicates. Anal Biochem 379, 127-129. Wilmore, H.P., McClive, P.J., Smith, C.A., Sinclair, A.H., 2005. Expression profile of the RNA- binding protein gene hermes during chicken embryonic development. Dev Dyn 233, 1045-1051. Wu, H., Sun, S., Tu, K., Gao, Y., Xie, B., Krainer, A.R., Zhu, J., 2010a. A splicing-independent function of SF2/ASF in microRNA processing. Mol Cell 38, 67-77. Wu, J., Li, C., Zhao, S., Mao, B., 2010b. Differential expression of the Brunol/CELF family genes during Xenopus laevis early development. Int J Dev Biol 54, 209-214. 215 Wu, J.I., Reed, R.B., Grabowski, P.J., Artzt, K., 2002. Function of quaking in myelination: regulation of alternative splicing. Proc Nat Acad Sci U S A 99, 4233-4238. Xiao, L., Cui, Y.H., Rao, J.N., Zou, T., Liu, L., Smith, A., Turner, D.J., Gorospe, M., Wang, J.Y., 2011. Regulation of cyclin-dependent kinase 4 translation through CUG-binding protein 1 and microRNA-222 by polyamines. Mol Biol Cell 22, 3055-3069. Xu, X., Yang, D., Ding, J.H., Wang, W., Chu, P.H., Dalton, N.D., Wang, H.Y., Bermingham, J.R., Jr., Ye, Z., Liu, F., Rosenfeld, M.G., Manley, J.L., Ross, J., Jr., Chen, J., Xiao, R.P., Cheng, H., Fu, X.D., 2005. ASF/SF2-regulated CaMKIIdelta alternative splicing temporally reprograms excitation-contraction coupling in cardiac muscle. Cell 120, 59-72. Xu, X.Q., Soo, S.Y., Sun, W., Zweigerdt, R., 2009. Global expression profile of highly enriched cardiomyocytes derived from human embryonic stem cells. Stem Cells 27, 2163- 2174. Yang, Y., Mahaffey, C.L., Berube, N., Maddatu, T.P., Cox, G.A., Frankel, W.N., 2007. Complex seizure disorder caused by Brunol4 deficiency in mice. PLoS Genet 3, e124. Yano, M., Hayakawa-Yano, Y., Mele, A., Darnell, R.B., 2010. Nova2 regulates neuronal migration through an RNA switch in disabled-1 signaling. Neuron 66, 848-858. Yatskievych, T.A., Ladd, A.N., Antin, P.B., 1997. Induction of cardiac myogenesis in avian pregastrula epiblast: the role of the hypoblast and activin. Development 124, 2561- 2570. Yeo, G.W., Xu, X., Liang, T.Y., Muotri, A.R., Carson, C.T., Coufal, N.G., Gage, F.H., 2007. Alternative splicing events identified in human embryonic stem cells and neural progenitors. PLoS Comput Biol 3, 1951-1967. Zaffran, S., Astier, M., Gratecos, D., Semeriva, M., 1997. The held out wings (how) Drosophila gene encodes a putative RNA-binding protein involved in the control of muscular and cardiac activity. Development 124, 2087-2098. Zhang, C., 2009. Novel functions for small RNA molecules. Curr Opin Mol Ther 11, 641-651. Zhang, C., Lee, K.Y., Swanson, M.S., Darnell, R.B., 2013. Prediction of clustered RNA-binding protein motif sites in the mammalian genome. Nuc Acids Res 41, 6793-6807. Zhang, C., Zhang, Z., Castle, J., Sun, S., Johnson, J., Krainer, A.R., Zhang, M.Q., 2008a. Defining the regulatory network of the tissue-specific splicing factors Fox-1 and Fox-2. Genes Dev 22, 2550-2563. Zhang, L., Lee, J.E., Wilusz, J., Wilusz, C.J., 2008b. The RNA-binding protein CUGBP1 regulates stability of tumor necrosis factor mRNA in muscle cells: implications for myotonic dystrophy. J Biol Chem 283, 22457-22463. Zhang, W., Haiying, L., Kyoungha, H., Grabowski, P., 2002. Region-specific alternative splicing in the nervous system: Implications for regulation by the RNA-binding protein NAPOR. RNA 8, 671-685. Zhang, X., Azhar, G., Furr, M.C., Zhong, Y., Wei, J.Y., 2003. Model of functional cardiac aging: young adult mice with mild overexpression of serum response factor. Am J Physiol Regul Integr Comp Physiol 285, R552-560. Zhou, Z., Fu, X.D., 2013. Regulation of splicing by SR proteins and SR protein-specific kinases. Chromosoma 122, 191-207. Zhu, B., Ramachandran, B., Gulick, T., 2005. Alternative pre-mRNA splicing governs expression of a conserved acidic transactivation domain in myocyte enhancer factor 2 factors of striated muscle and brain. J Biol Chem 280, 28749-28760. 216