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An internal promoter underlies the difference in disease severity between N- and C-terminal truncation mutations of Titin in zebrafish

  1. Jun Zou
  2. Diana Tran
  3. Mai Baalbaki
  4. Ling Fung Tang
  5. Annie Poon
  6. Angelo Pelonero
  7. Erron W Titus
  8. Christiana Yuan
  9. Chenxu Shi
  10. Shruthi Patchava
  11. Elizabeth Halper
  12. Jasmine Garg
  13. Irina Movsesyan
  14. Chaoying Yin
  15. Roland Wu
  16. Lisa D Wilsbacher
  17. Jiandong Liu
  18. Ronald L Hager
  19. Shaun R Coughlin
  20. Martin Jinek
  21. Clive R Pullinger
  22. John P Kane
  23. Daniel O Hart
  24. Pui-Yan Kwok
  25. Rahul C Deo  Is a corresponding author
  1. University of California, San Francisco, United States
  2. University of North Carolina at Chapel Hill, United States
  3. Brigham Young University, United States
  4. University of Zurich, Switzerland
  5. University of California, United States
  6. California Institute for Quantitative Biosciences, United States
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Cite this article as: eLife 2015;4:e09406 doi: 10.7554/eLife.09406

Abstract

Truncating mutations in the giant sarcomeric protein Titin result in dilated cardiomyopathy and skeletal myopathy. The most severely affected dilated cardiomyopathy patients harbor Titin truncations in the C-terminal two-thirds of the protein, suggesting that mutation position might influence disease mechanism. Using CRISPR/Cas9 technology, we generated six zebrafish lines with Titin truncations in the N-terminal and C-terminal regions. Although all exons were constitutive, C-terminal mutations caused severe myopathy whereas N-terminal mutations demonstrated mild phenotypes. Surprisingly, neither mutation type acted as a dominant negative. Instead, we found a conserved internal promoter at the precise position where divergence in disease severity occurs, with the resulting protein product partially rescuing N-terminal truncations. In addition to its clinical implications, our work may shed light on a long-standing mystery regarding the architecture of the sarcomere.

https://doi.org/10.7554/eLife.09406.001

eLife digest

The heart is able to beat partly because of a large protein called Titin that helps to give heart muscle its elasticity. Mutations that shorten the gene that encodes Titin can cause part of the heart to become enlarged and weakened, a condition called dilated cardiomyopathy. Some people with shortened copies of this protein have a mild form of cardiomyopathy and are able to lead relatively normal lives. Others develop more severe symptoms that prevent the heart from pumping blood effectively and may even cause the individual to need a heart transplant.

Genetic studies have revealed that mutations that shorten the Titin protein by disrupting the portion of the gene corresponding to the latter two-thirds of the protein (which encodes the so-called “C-terminal” end of the protein) cause more severe symptoms than mutations that occur near the start of the gene. But it is not clear why the location of the mutation matters.

To investigate this problem, Zou et al. used a gene-editing tool called CRISPR to create genetically engineered zebrafish. These fish had mutations at one of six different points in the gene that encodes the zebrafish version of Titin. Just as with humans, mutations near the C-terminal end of the gene caused more severe muscle problems in the fish.

Specifically, Zou et al. found that the worst disease was associated with mutations that occurred at or after a “promoter” region within the gene and near this C-terminal end. Normally, the promoter produces an independent smaller form of the Titin protein, which helps to reduce the severity of muscle problems in zebrafish that have mutations near the start of the gene. However, mutations near the C-terminal end of the gene also damage this smaller form, preventing this failsafe from working, and so lead to more severe symptoms. Zou et al. also found this promoter to be active in both mouse and human hearts.

Future work will focus on learning how this smaller form of Titin works to help muscle develop and withstand stress and determine whether increasing its production can overcome the more severe forms of disease.

https://doi.org/10.7554/eLife.09406.002

Introduction

The use of genetics in clinical medicine depends on knowledge that an identified mutation confers risk of disease. Recent guidance issued by the American College of Medical Genetics emphasizes that the strongest form of evidence supporting causality of a mutation is whether it results in a truncation of the protein (nonsense, frameshift or canonical splice-site mutations), specifically for genes where loss-of-function mutations are known to cause disease (Richards et al., 2015). Truncating mutations within alternatively spliced exons and at the extreme C-terminus of the protein must be interpreted with caution, as they may have little to no impact on protein function. However, the guidelines do not address whether truncating mutations at different positions along the length of the protein might differ in phenotype severity or mechanism of action, a finding that could complicate interpretation of truncation mutations for a broad range of inherited diseases.

Truncations in the giant sarcomeric protein Titin (TTN) reveal patterns that are at odds with this uniform assignment of causality. TTN lies along the length of the sarcomere and is thought to mediate passive tension of the muscle fiber, while also serving as a binding scaffold for a large number of proteins, including actin and myosin (Lewinter and Granzier, 2013). Individual regions of TTN are annotated according to their localization to the Z-disc, I-band, A-band, and M-line regions of the sarcomere visualized on immuno-electron micrographs Fürst et al., 1988. Truncating and/or missense mutations in TTN result in cardiac (dilated cardiomyopathy [DCM]) and skeletal muscle (myofibrillar myopathy) disease (Chauveau et al., 2014). Recent estimates suggest that truncation mutations in TTN may explain as much as 25% of DCM (Herman et al., 2012), a disease which affects upwards of 1 in 2500 individuals (Codd et al., 1989). Although truncations are found along the length of TTN in DCM patients, in patients with the most severe forms of DCM, truncations reside within the C-terminal two-thirds of the protein (from amino acid 14,760 onwards, Figure 1A), specifically in the distal I-band and A-band (we will refer to these as C-terminal truncations) (Herman et al., 2012; Roberts et al., 2015). Multiple TTN exons are alternatively spliced (Bang et al., 2001), with many having little to no inclusion within cardiac isoforms, and splicing has been proposed to at least partially explain the exclusion of N-terminal mutations in patients with end-stage DCM (Herman et al., 2012). Nonetheless, over 5500 amino acids in the N-terminal region of the protein are constitutive (percent spliced in or PSI >0.9, Figure 1B), implying the importance of other mechanisms.

Figure 1 with 1 supplement see all
Generation of six stable zebrafish lines with truncating mutations in ttna, the zebrafish orthologue of TTN, using CRISPR/Cas9 technology.

(A) Schematic of the TTN meta-transcript (Ensembl ID ENST00000589042), including all known exons with the exception of the Novex-3 exon, which is exclusive to the Novex-3 isoform (Ensembl ID ENST00000360870). Sarcomeric regions are denoted by different colors. A heatmap of PSI values computed from RNA-Seq data of two DCM hearts is shown below the cartoon representation of the transcript. Above the schematic, amino acid positions of truncating mutations of TTN in end stage dilated cardiomyopathy identified in a prior study (Roberts et al., 2015) are highlighted (gold squares). We restricted our representation to nonsense, frameshift or canonical splice-site mutations and did not include predicted disruptions of putative splice enhancers. A truncating mutation mapping to the Novex-3 exon is not shown. Below the schematic, the corresponding human amino acid positions of zebrafish exons disrupted through CRISPR/Cas9 technology in our work are depicted, including six stable mutant lines (n1, n2, n3, c1, c2, c3, gold diamonds) and individual exons targeted by high dose guide RNA/Cas9 injection (gold triangles). (B) Many of the exons not mutated in end-stage DCM are constitutive (i.e. show minimal alternative splicing). Histogram representing the PSI frequency distribution for TTN amino acids N-terminal to amino acid 14,760, which is the most N-terminal amino acid in which a definitive truncation mutation is found in an end-stage DCM patient. The median PSI value was computed for each TTN exon using RNA-Seq datasets from hearts of two patients with DCM (GEO Accession GSM1380722, GSM1380723).

https://doi.org/10.7554/eLife.09406.003

Results

We sought to address the basis of this variation of severity with mutation position. For a disease such as DCM, in which inheritance is most commonly observed as adult-onset autosomal dominant with variable penetrance, deciphering such mechanisms in mouse models can be challenging, as heterozygotes tend to show minimal phenotypes and homozygotes tend to be embryonic lethal (Gramlich et al., 2009). We selected the vertebrate zebrafish model for several reasons. Firstly, zebrafish has high sequence conservation of sarcomeric proteins with mammals and recapitulate disease phenotypes with loss-of-function of multiple established Mendelian cardiomyopathy genes (Dahme et al., 2009). In fact, forward genetic screens have generated zebrafish Titin mutants, although in only one case is the position of the mutation known (Xu et al., 2002; Steffen et al., 2007; Myhre et al., 2014). Secondly, zebrafish can live for up to two weeks with a non-contractile heart, thus allowing mechanistic studies of homozygote mutants. Thirdly, the relatively low cost and large clutch size of zebrafish allow the ability to generate multiple mutant lines and provide statistical power for robust conclusions.

As a result of an ancestral genome duplication event, zebrafish include two titin genes: ttna and ttnb (Xu et al., 2002; Steffen et al., 2007; Seeley et al., 2007). Ttna shares considerable similarity with the human TTN gene (Figure 1—figure supplement 1, showing ‘meta-transcripts’ with all possible exons included). The zebrafish and human proteins are similar in length (∼32,000–35,000 amino acids depending on the splice isoform) and share 56% identity and 82% homology. They have a nearly identical domain organization, with three immunoglobulin-like of domains (proximal, mid, and distal), an elastic PEVK region, N2A and N2B domains which distinguish cardiac and skeletal isoforms, a fibronectin-immunoglobulin repeat region that mediates binding to myosin, and a C-terminal kinase domain. In both species, alternative splicing targets the middle Ig-like and PEVK domains, which modulate passive tensile properties of the sarcomere (Granzier and Labeit, 2004). Cardiac isoforms typically include much of this elastic region as well as the N2B ± N2A domains, while skeletal isoforms include the N2A domain but exclude N2B as well as much of the elastic region. Multiple mutants isolated from forward genetic screens coupled with morpholino knockdown analysis have demonstrated that ttna is essential for cardiac sarcomere formation in zebrafish (Xu et al., 2002; Myhre et al., 2014; Seeley et al., 2007).

Although the paralagous zebrafish ttnb gene is dispensable for cardiac sarcomere development (Steffen et al., 2007; Seeley et al., 2007), it plays an essential role in skeletal muscle sarcomerogenesis. It is expressed at comparable levels to ttna in skeletal muscle (as opposed to ∼2/3 of ttna abundance in heart). In contrast to human TTN or zebrafish ttna, the ttnb protein has abbreviated versions of the middle Ig-like and PEVK elastic domains (Figure 1—figure supplement 1) and thus more closely resembles skeletal rather than cardiac muscle isoforms of ttna. Morpholino-based skipping of the N2A exon of either ttna or ttnb results in severe disruption of skeletal muscle sarcomeric architecture, highlighting the mutual contribution of the two genes (Seeley et al., 2007).

Using CRISPR/Cas9 technology (Jinek et al., 2012), we engineered six truncating mutations within the major cardiac TTN orthologue, ttna, (Figure 1A, Table 1), so that the resulting protein product would be truncated at either the Z-disk (n1), proximal or mid I-band (n2 and n3) or proximal, mid, or distal A-band (c1, c2, and c3). From wild-type (WT) zebrafish embryo transcriptome data, we anticipated that all exons would be constitutive in both cardiac and skeletal muscle (PSI >0.99, Table 1). Mutant lines were bred for two generations to minimize any off-target effects from the genome editing, and we studied the F3 generation for cardiac and skeletal muscle consequences.

Table 1

Characteristics of six stable zebrafish truncation mutant lines. Size of insertion/deletion, PSI of corresponding exon in embryonic zebrafish heart and skeletal muscle, protein change using standard nomenclature (23), and corresponding human amino acid protein for the TTN meta-transcript are shown.

https://doi.org/10.7554/eLife.09406.005
Mutant IDMutation typePSI of mutated exon (heart/skeletal muscle)Zebrafish protein change#Orthologous human amino acid position*
n17bp insertion1.000/1.000p.E1463fsX31697
n21bp insertion1.000/1.000p.A2815GfsX33048
n32bp deletion1.000/1.000p.Q7597GfsX89693
c12bp deletion0.996/1.000p.E15193GfsX817996
c24bp deletion1.000/0.995p.T18311GfsX821215
c38bp insertion1.000/0.996p.A29052RfsX832003
  1. #Amino acid numbering is relative to ENSDART00000109099. *Amino acid numbering is relative to TTN meta-transcript.

As expected (Xu et al., 2002; Gramlich et al., 2015), ttna truncations showed no gross phenotype in heterozygotes, with all heterozygotes attaining sexual maturity. We thus focused on homozygote mutants, with the expectation that partial rescue from ttnb could potentially elicit phenotypic differences among mutants, especially in skeletal muscle where it is expressed at comparable levels to ttna and has a documented essential role. N-terminal (ttnan1/n1, ttnan2/n2, ttnan3/n3, collectively referred to as ttnan/n) and C-terminal (ttnac/c) truncations all showed severe deficits in cardiac contractility (Video 1) and typically died within two weeks of birth. Immunostaining revealed severe cardiac sarcomeric disarray (Figure 2A).

C-terminal ttna truncations result in a severe skeletal muscle phenotype while N-terminal truncations are indistinguishable from wild-type.

Fixed heart (A) and skeletal muscle (B) samples of 72 hpf ttnawt/wt, ttnan/n and ttnac/c mutant embryos were analyzed by immunostaining for α–actinin, which highlights Z-disc architecture. The cardiac sarcomere was disarrayed in all mutants. However, in skeletal muscle ttnac/c mutants demonstrated severe sarcomeric disarray while ttnan/n mutants retained sarcomeric architecture. Scale bar: 10 uμm. (C) All targeted TTN exons are constitutive (i.e. not alternatively spliced) in both cardiac and skeletal muscle. PSI values computed for each mutated exon using RNA-Seq data for dissected hearts and trunk skeletal muscle for various mutant genotypes at 72 hpf. Wild-type fish were analyzed at both 72 hpf and adulthood. Analysis was limited to samples with a sufficient number of exon-exon junction reads to accurately estimate PSI (Pervouchine et al., 2013). (D) Nonsense-mediated decay reduces mutant transcript levels to ∼20–25% of wild-type, but does not vary substantially across mutations. Targeted RNA-Seq was used to determine the ratio of reads derived from the mutant allele vs. the wild-type allele in ttnan/wtand ttnac/wt heterozygote mutants, which serves as an estimate of nonsense-mediated decay efficiency.

https://doi.org/10.7554/eLife.09406.006
Video 1
Cardiac contraction for wild-type, ttnan/n and ttnac/c mutants.

All embryos were imaged at 72 hpf.

https://doi.org/10.7554/eLife.09406.007

In contrast to their cardiac phenotypes, ttna mutants demonstrated a sharp division in development of skeletal muscle disease. All three ttnan/n mutants were capable of grossly normal motion while the ttnac/c truncations were entirely unable to move (Video 2). Immunostaining supported the distinction, with ttnac/c mutants having highly distorted sarcomeric architecture while ttnan/n architecture closely resembled wild-type (Figure 2B). To confirm transcription of all mutant isoforms, assess nonsense-mediated decay, and rule out selective alternative splicing of ttnan/n mutant exons, we performed RNA-Seq of heart (cardiac muscle) and trunk (skeletal muscle) of 3 day old embryos, as well as adult fish. All mutated exons appeared constitutive in both heart and skeletal muscle, across all genotypes (Figure 2C). We next examined the extent of nonsense-mediated decay (NMD) for each mutant allele by performing targeted RNA sequencing of heterozygote embryos and comparing the number of reads for wild-type and mutant exons. In keeping with typical estimates for NMD efficiency (Zetoune et al., 2008), we saw ∼75–80% of mutant transcript being degraded (Figure 2D). However, no substantial variation was seen across the mutants.

Video 2
Motility assessment for wild-type, ttnan/n and ttnac/c mutants.

All embryos were imaged at 72 hpf.

https://doi.org/10.7554/eLife.09406.008

Given that variability in alternative splicing or NMD could not explain the selective skeletal muscle disarray in ttnac/c mutants, we hypothesized that this variation in severity could be due to ttnac/c mutants interfering with the action of ttnb, although the ratio of mutant to wild-type transcript of ∼20% made this an unlikely scenario. Knockdown of ttnb by morpholino disrupted ttnan/n skeletal muscle morphology indicating that the intact skeletal muscle sarcomeric architecture in ttnan/n mutants arose at least in part from rescue by ttnb protein (Figure 3A), although the phenotype was not nearly as severe as that of ttnac/c mutants. To investigate a dominant negative mechanism for ttnac/c mutants, we performed a morpholino knockdown of ttna, targeting a splice site of an N-terminal constitutive exon (exon 4), which would be expected to introduce an N-terminal frameshift on top of the same C-terminal mutant allele. If the differences between ttnan/n and ttnac/c mutants arose simply from inhibitory effects of the ttnac/c truncated product, then introducing an upstream frameshift should disrupt production of the dominant negative protein product even further and restore sarcomeric architecture. Knockdown by morpholino was effective (∼78% knockdown efficiency, data not shown) and the combined action of NMD and morpholino knockdown would be expected to reduce mutant ttna protein to only ∼5% of the level of ttnb protein. Nonetheless we were unable to rescue skeletal muscle architecture in ttnac/c mutants (Figure 3B). We thus concluded that ttnac/c mutants do not act as dominant negatives and that an alternative explanation was needed.

C-terminal ttna truncations do not act as dominant negatives.

(A) Knockdown of ttnb by morpholino injection worsens skeletal muscle sarcomeric architecture in ttnan/n mutants. Fixed skeletal muscle samples of 72 hpf ttnan1/n1 mutant embryos were analyzed by immunostaining for α–actinin. (right) Cartoon representation of ttna and ttnb proteins, with a premature N-terminal stop codon in ttna (gold star). The purple lines indicate morpholino disruption of the ttnb transcript. Scale bar: 10 uμm. (B) Knockdown of ttna by morpholino injection does not rescue skeletal muscle architecture in ttnac1/c1 mutants. ttnac1/c1 embyros were injected with a ttna splice-site morpholino to the exon 4-intron 4 junction at the 1–2 cell stage and embryos were examined at 72 hpf. At this morpholino dose, knockdown efficacy was estimated at close to 80% and nearly complete cessation of cardiac contraction was achieved in >90% of wild-type embryos (data not shown). Immunostaining for α–actinin revealed no improvement in skeletal muscle architecture. (right) Cartoon representation of mutant ttna proteins, with a premature C-terminal stop codon (gold star). The purple lines indicate disruption of the ttna transcript using a morpholino that is expected to introduce an N-terminal truncation upstream of the C-terminal truncation.

https://doi.org/10.7554/eLife.09406.009

Another possible explanation for the discrepancy in disease severity between ttnan/n and ttnac/c mutants could be an internal promoter that produces a C-terminal isoform, which could partially rescue N-terminal truncations, especially in conjunction with expression of ttnb. Even though N- and C-terminal truncating mutations would both disrupt full-length ttna, the latter would also disrupt a C-terminal isoform while the former would not. Interestingly invertebrates, including C. elegans and D. melanogaster, have distinct genes that encode for protein products corresponding to the N-terminal (Z-disc and proximal I-band) or C-terminal (distal I-band and A-band and M-line) portions of Titin (Bullard et al., 2002). The Novex-3 isoform in humans (discussed below) does in fact include only the Z-disc and proximal I-band portions of TTN, but no isoform corresponding only to the C-terminal portion of TTN has ever been reported in vertebrates.

Using the exon boundaries defined by ttnan3/n3 and ttnac1/c1, we examined zebrafish and mouse cardiac and skeletal muscle transcriptomic data for patterns of reads indicating a promoter upstream of one of the ttna exons. We noticed an accumulation of reads 5’ to exon 116 (e116, ENSDART00000109099) in zebrafish skeletal muscle (Figure 4A) and in the corresponding exon of mouse heart (e223, ENSMUST00000099981, Figure 4A). We hypothesized that these reads arose from transcription from an alternative promoter within the e115-e116 intron. 5’-RACE confirmed an alternative transcription start site (TSS), 110 bp upstream of e116 (within the e115-e116 intron) in zebrafish skeletal muscle, with an in-frame initiator methionine located 7 amino acids upstream of e116. To establish conservation in mammals, we repeated 5’-RACE on adult mouse and fetal human cardiac tissue and found a TSS in the upstream intron (257bp upstream of e223 in mouse; 256bp upstream of the orthologous exon, e240, in humans), with a putative initiator methionine 12 amino acids upstream of the relevant exon. Using PCR, we confirmed the presence of a multi-exonic transcript that includes this alternative promoter site in zebrafish, mouse, and human cardiac tissue (Figure 4B).

Figure 4 with 2 supplements see all
An internal promoter at the distal I-band explains phenotypic differences between N- an C-terminal ttna mutants.

(A) RNA-Seq data from 72 hpf zebrafish trunk (left) and E12.5 mouse hearts (right) depicting accumulation of reads within the intron, upstream of the orthologous exons 116 (zebrafish) and 223 (mouse). The Ttn gene is on the negative strand and so reads within the upstream intron are shown to the right of the black bar, which indicates the position of the relevant exon. (B) Transcription from an alternative Titin promoter occurs in zebrafish, mouse, and human heart. (left) PCR amplification using a primer in the upstream exon or an internal primer at or near the internal TSS (as determined by 5’-RACE) as forward primer and a shared reverse primer was performed using cDNA from zebrafish, mouse and human heart. For all three species, products of the expected size were found, supporting transcription from an internal promoter. (right) Cartoon representation of PCR amplification scheme, with exon numbering according to zebrafish transcript. The zebrafish 5’ UTR is shorter than that of mouse and human. (C) (left) H3K4me3 ChIP-Seq data from human fetal leg muscle (GEO Accession GSM1058781), indicating an active promoter overlies exon 240, which is orthologous to exon 116 in zebrafish. The peak at the far right of the figure represents the conventional human TTN promoter. (right) DNAse-Seq data from human fetal heart (GEO Accession GSM665830) indicates highly accessible chromatin overlying exon 240, which typically demarcate enhancers or promoters (Neph et al., 2013). The accessible chromatin peak at the far right is the conventional TTN promoter. The gold triangle in both panels indicates the genomic position of the human internal promoter TSS identified through 5’-RACE. (D) Phenotypic divergence in skeletal myopathy in ttna truncations occurs at the exons flanking the alternative promoter. Cas9 protein and high dose gRNAs corresponding to exon 115 or exon 116 were injected into 1 cell embryos. At 72 hpf, embryos with complete or near complete cessation of cardiac contraction were collected and immunostained for α–actinin to assess skeletal muscle architecture. (E) An ATG morpholino targeting the intronic region upstream of the novel initiator methionine is sufficient to create skeletal muscle disarray on a ttnan3/n3 background. 1–2 cell embryos from a ttnan3/wt cross were injected with an intronic morpholino and ttnan3/n3 homozygotes immunostained for α–actinin to assess sarcomere architecture in the skeletal muscle. No skeletal muscle disarray was seen in wild-type fish injected with the internal ATG morpholino (data not shown).

https://doi.org/10.7554/eLife.09406.010

We next consulted publicly available chromatin accessibility and activation data to find supporting evidence for an internal TTN promoter at this site. DNAse-Seq is a powerful tool to identify regions of accessible chromatin, which typically correspond to promoter or enhancer regions (Neph et al., 2013). Examination of multiple fetal human cardiac and skeletal muscle DNA-Seq datasets revealed a prominent peak overlying e240, the orthologous exon to e116 in humans (Figure 4C, Figure 4—figure supplement 1). A similar peak was found in H3K4me3 Chip-Seq data, a mark for active promoters (Zhou et al., 2010), from skeletal muscle and cardiac tissue (Figure 4C, Figure 4—figure supplement 1). Unbiased epigenetic data thus provides additional strong support for an internal promoter. For convenience, and as a counterpart to the N-terminal Novex-3 isoform, we hereafter refer to this C-terminal Titin isoform as the Cronos isoform.

If a protein product arising from the ttna internal promoter ameliorates disease severity in ttnan/n but not ttnac/c mutants, we should be able to pinpoint the phenotypic transition at e116 – i.e. truncation mutants N-terminal to e116 should have mild phenotypes while those at or C-terminal to e116 should have severe phenotypes. Interestingly, the corresponding human genomic position (the e239-e240 intron in ENST00000589042) represents the exact transition point for where disease severity markedly increases in DCM TTN truncation patients (Figure 1A). We used high dose guide RNA (gRNA) and Cas9 protein injections to exons 113, 114, 115, 116, and 122 and examined cardiac and skeletal muscle architecture and function in the resulting fish, focusing on embryos with nearly complete cessation of cardiac beating, as these would be expected to have a high likelihood of homozygous genomic disruption (given that only homozygote and not heterozygote ttnan/n or ttnac/c mutants have a cardiac phenotype). As expected, e113, e114 and e115 F0 mutants showed intact skeletal muscle architecture while e116 and e122 mutants had severe skeletal muscle disruption (Figure 4D, Figure 4—figure supplement 2). To confirm that the Cronos isoform protein product was actually translated, we used agarose protein electrophoresis and found that a higher mobility (smaller size) protein product was indeed found in wild-type and ttnan/n mutants but not in ttnac/c mutants (Figure 5A). Finally, a translation start site morpholino targeted upstream of the actual alternative initiator methionine (and 38 nucleotides upstream of the intron-exon junction) was sufficient to completely impair motility and disrupt skeletal muscle architecture in the ttnan3/n3 mutant (Figure 4E, Video 3), further supporting the hypothesis that the Cronos product can rescue skeletal muscle architecture in ttnan/n mutants. No obvious impact of the morpholino on alternative splicing of neighboring exons was seen (data not shown).

Premature termination codons in ttna differentially affect the levels of Cronos and full-length ttna depending on mutation location.

(A) 1.0% agarose gel reveals higher molecular weight bands corresponding to different splice isoforms of ttnb (all fish) and ttna (wild-type only). The arrow represents Cronos isoform of ttna, which is absent in C-terminal truncation mutants and in zebrafish treated with an ATG morpholino against this isoform. Ttna and ttnb expression results in a complex distribution of splice isoforms, whose identity and composition vary over early development (Steffen et al., 2007). (B) NMD results in a high ratio of Cronos to full-length ttna transcript levels in N-terminal but not C-terminal truncation mutants. Quantitative real-time PCR was used to estimate relative levels of the Cronos and full-length ttna trancripts for wild-type and mutant embryos. No efficiency difference was seen for the corresponding primer pairs.

https://doi.org/10.7554/eLife.09406.013
Video 3
Motility for wild-type and ttnan/n mutants injected with internal promoter ATG morpholino
https://doi.org/10.7554/eLife.09406.014

Given that NMD substantially reduces the levels of ttna transcripts with premature termination codons (Figure 2D), we reasoned that we should be able to distinguish N- and C-terminal truncation mutants by the ratio of Cronos to full-length transcript. In ttnan/n mutants, which would degrade full-length ttna but not Cronos, we would anticipate a higher Cronos:full-length ratio than in ttnac/c mutants mutants, which should degrade both transcripts approximately equally. As expected, we see a much higher ratio of Cronos:full-length transcript levels in ttnan/n mutants than in ttnac/c mutants (Figure 5B).

We next surveyed the expression levels of Cronos in cardiac and skeletal muscle in zebrafish at different developmental stages using both in situ hybridization (Figure 6A) and real-time PCR (Figure 6B). Cronos is expressed at high levels in zebrafish skeletal muscle during development (∼2:1 ratio of Cronos to full-length ttna), but drops off sharply during adulthood (∼1:70 ratio). Although the Cronos expression in zebrafish heart also appears to be developmentally regulated, it is much lower than in skeletal muscle, with a ∼1:5 ratio at 72 hpf and 1:30 ratio in adulthood. This low cardiac Cronos expression (coupled with a minor role of ttnb in zebrafish hearts) is a likely reason for the severe and indistinguishable cardiac sarcomere disruption in ttnan/n and ttnac/c mutants.

Tissue and developmental profile of Cronos and ttna expression in zebrafish and mouse heart and skeletal muscle.

(A) In situ hybridization at 72 hrpf (left) and 24 and 48 hpf (right) reveal prominent expression of both Cronos and full length ttna isoforms in zebrafish somites. In contrast, Cronos expression in developing zebrafish heart (arrow) is at low levels. (B) Quantitative PCR estimates of the ratio of Cronos to full-length ttna reveals prominent expression in developing zebrafish skeletal muscle, but markedly reduced expression in developing heart and in adult heart and skeletal muscle. (C) Quantitative PCR estimates of the ratio of Cronos to full-length Ttn reveals prominent expression in developing mouse hearts, which diminishes through development. Skeletal muscle Cronos expression is comparable in early postnatal mouse, but diminished in adulthood and varies by skeletal muscle bed. EDL = extensor digitorum longus. LV/RV = left/right ventricle.

https://doi.org/10.7554/eLife.09406.015

Given that the primary observation that motivated this work was variation in phenotypic severity in cardiomyopathy patients, we next looked to see if there was appreciable Cronos expression in mammalian hearts. We found that Cronos levels were much higher in developing mouse hearts than in zebrafish, with nearly equal transcript levels of Cronos and full-length Ttn (Figure 6C) at embryonic day 12.5. Thereafter mouse cardiac expression of Cronos decreases to ∼20–30% of full-length Titin. Skeletal muscle expression varies across development and in different muscle beds with a 1:5 ratio in mouse hindlimb at P2, and between 1:16 and ∼1:1000 ratios in adult extensor digitorum longus (fast-twitch) and soleus (slow-twitch) muscle, respectively.

Although our zebrafish work explains why a milder phenotype would be expected in individuals with truncation mutants upstream of the alternate promoter, it does not answer whether such N-terminal truncations would still result in deleterious manifestations in heterozygote patients. Specifically, an open question remains whether all TTN truncations lead to cardiac or skeletal muscle phenotypes, or conversely, do any TTN truncations lack cardiac or skeletal muscle consequences. In a disease such as dilated cardiomyopathy, with a variable age of onset, it can be challenging to derive general conclusions from observational data of control subjects, many of whom are young or middle-aged. Furthermore, self-reports of ‘health’ may not reveal limitations in an individual’s ability to attain full exercise potential, especially with aging.

To assess whether any TTN truncations are compatible with a high degree of cardiovascular fitness throughout life, we turned to a population genetics approach, and sequenced the complete TTN exome (as well as that of 104 other Mendelian cardiac disease genes, Supplementary file 1A) in 199 competitive senior athletes who entered as contestants in the Huntsman World Senior Games (median age 73, Table 2). We compared the frequency and distribution of TTN truncation mutations to those found in unselected controls and healthy volunteers from a recent sequencing study (CTLlit) (Roberts et al., 2015). The overall rate of TTN truncations did not differ between senior athletes and CTLlit (1.5 vs 1.2%, p = 0.91). However, the distribution of TTN truncations differed significantly between groups (Figure 7, Figure 7—figure supplement 1): TTN truncations in senior athletes mapped exclusively to the extreme C-terminal exon of the rare Novex-3 isoform (Bang et al., 2001), at a nearly 8-fold higher rate than CTLlit (p = 0.003). This exon is found only in the Novex-3 isoform, which is expressed at low levels in the human heart (<10% of the levels of major TTN isoforms [Roberts et al., 2015]), and we would thus expect truncations in this exon would have little effect on TTN function. We next analyzed the ratio of mutants up- and downstream of Cronos in affected DCM patients, literature controls and senior athletes (Figure 7). As expected, in end-stage DCM we observed a much higher rate of mutants C-terminal to Cronos than N-terminal (30:1), followed by unselected DCM (37:11), literature controls (6:11, 2:2, 10:7 for FHS, healthy volunteers and JHS) and senior athletes (0:3).

Table 2

Demographic characteristics of 199 senior athletes genotyped by capture-based targeted sequencing for this study. 

https://doi.org/10.7554/eLife.09406.016
Number199
Age73 (IQR 68-77)
Sex60% Male
Race97% European American
SportsTrack and field, volleyball, cycling, softball, marathon, …
Figure 7 with 1 supplement see all
TTN truncations are found preferentially upstream of the Cronos promoter in controls but downstream of Cronos in DCM cases.

Ratio of the number of truncating mutations found C-terminal to the position of the Cronos promoter to those found N-terminal, in DCM cases and controls derived from the literature (Roberts et al., 2015) and in senior athletes.

https://doi.org/10.7554/eLife.09406.017

Discussion

Loss of function experiments across a number of systems have demonstrated Titin’s importance in diverse aspects of sarcomere development and function, including the ordered assembly of sarcomeric constituents (Xu et al., 2002; van der Ven et al., 2000), cardiomyocyte systolic (Hinson et al., 2015) and diastolic function (Radke et al., 2007), and resistance to neurohormonal and vasoconstrictor stresses (Gramlich et al., 2009). Our work further contributes the discovery of a conserved internal promoter in TTN, whose protein product is essential for skeletal muscle sarcomere development in zebrafish. The Cronos promoter is prominently expressed in developing mouse heart, and its location coincides strikingly with the position of TTN truncations in patients with the most severe forms of dilated cardiomyopathy. Our working model is that disruption of both the full-length TTN and Cronos protein products results in more severe disease than disruption of the full-length product alone.

Throughout this work, we have focused on skeletal muscle architecture and function in zebrafish ttna homozygote mutants, given that this tissue demonstrates a sharp and rapidly developing phenotypic distinction between N- and C-terminal Titin truncations. The early onset of this unambiguous phenotypic difference combined with the ease of genetic and transcriptomic manipulation using CRISPR/Cas9 and morpholino technology allowed efficient mapping of the novel Cronos internal promoter as well as testing of alternative mechanistic hypotheses for this phenomenon. Nonetheless, implicit in our work is the assumption that findings from zebrafish skeletal muscle sarcomere development in ttna homozygote mutants (albeit with the paralogous ttnb present for rescue) are relevant towards understanding disease development in humans with TTN mutations. Prominent differences between these systems must be noted. With the exception of certain childhood-onset forms of skeletal and cardiac myopathy arising from homozygous TTN truncations (Carmignac et al., 2007; Ceyhan-Birsoy et al., 2013), the mode of inheritance in TTN truncation mutation patients is heterozygous with marked variable expressivity and incomplete penetrance (Herman et al., 2012; Roberts et al., 2015; Itoh-Satoh et al., 2002; Norton et al., 2013; van Spaendonck-Zwarts et al., 2014). Moreover, disease onset usually occurs in adulthood. Such an inheritance pattern is typical of many of the familial forms of cardiomyopathy, and is believed to imply additional modifying effects, potentially from diverse sources as autoimmunity and viral infection as well as contributions from additional genetic variants (Mrh et al., 2015). Nonetheless, taking all of our results together, we find it a highly plausible explanation that a superimposed deficiency in expression of the Cronos protein product would result in the more severe forms of human cardiac disease seen in DCM patients with C-terminal TTN truncations.

With these limitations noted, our results have two clinical implications for titinopathy patients. Firstly, by precisely mapping the location of the alternate promoter to the e239-e240 intron in humans, we are in a better position to give mutation-specific guidance to TTN truncation patients regarding the potential severity of their disease, with the caveat that additional factors (genetic and environmental) also modulate phenotypic severity. Along these lines, given the overlapping distribution of mutations in unselected cases and controls, it is unlikely that highly accurate patient prognoses would be realized from genotype information alone (Figure 7—figure supplement 1), although there does appear to be nearly complete separation of mutation distribution for the more extreme phenotypes (end-stage DCM and senior athletes). Secondly, our work implies that strategies that augment activity of the internal TTN promoter and levels of the Cronos isoform might provide therapeutic benefit in TTN truncation patients, although it is unclear whether there is a restricted developmental window where this might be possible.

In addition to clinical ramifications, our work may shed light on a long-standing mystery regarding sarcomeric architecture. Electron microscopy reconstructions of serial sections of skeletal muscle dating back over 50 years presented a puzzle: the thin/thick filament arrangement in the sarcomere alters its geometry, progressing from a tetragonal lattice arrangement at the Z-band to a trigonal arrangement in the A-band (Pringle, 1968), with the transition taking place near the I-A junction (Traeger et al., 1983). The implication for TTN would be that there would be a resulting increase in the copies of TTN protein, with four at the Z-band and six at the A-band and M-line (Liversage et al., 2001), which is difficult to reconcile with the fact that a single TTN molecule spans the entire length, from Z-band to M-line. Our data provides a simple explanation for this observation, with the Cronos protein product selectively increasing copies of TTN at the distal I-band, A-band and M-line. Our work also proposes an alternative explanation for the consistently observed ‘T2’ band on protein gels, which migrates at the expected molecular weight of Cronos. This has invariably been labeled a C-terminal degradation product, as it reacts with A-band antibodies (Opitz et al., 2004). However, in keeping with Cronos expression patterns in development, the T2 band was prominently seen in developing mouse (Lahmers et al., 2004) and rat hearts (Opitz et al., 2004), but was markedly reduced in adulthood. It is possible that, at least in some situations, T2 and Cronos represent the same isoform.

These results also raise a number of questions related to the function of the Cronos isoform. Given its more marked expression in early development, is it important only for the initial sarcomerogenesis or in subsequent aspects of muscle function? Is it likely to be reactivated under stress? And how fast does it turn over compared to full-length TTN? Models with selective disruption and tagging of this isoform should begin to answer these questions. Finally, it is possible that unrecognized internal promoters influence disease severity in truncating mutations of other genes. It is likely a more nuanced diagnostic and prognostic interpretation of truncating mutations will be needed.

Materials and methods

sgRNA design, cloning, and in vitro transcription

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sgRNA’s were designed using an in-house Python script, which scans genomic sequences for 23 bp sequences with a terminal NGG (PAM sequence), and prioritizes a GG at the 5’ end because of the use of T7 RNA polymerase for gRNA synthesis. All sgRNA’s had only a single genome match. Oligonucleotides corresponding to the sgRNA sequence were annealed and then cloned into vector DR274 (Addgene plasmid 42250) as previously described (Hwang et al., 2013). For sgRNA synthesis, 100 ng of plasmid template was used in a T7 in vitro transcription reaction (MAXIscript T7, Life Technologies, Carlsbad, CA, AM1312M). RNA products were digested with DNAse I and purified by column (RNA Clean and Concentrator, Zymo Research, Irvine, CA, R1016).

Cas9 induced genomic cleavage and genotyping

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Recombinant Cas9 was prepared as previously described (Jinek et al., 2012). A mixture of recombinant Cas9 (0.5 ug/uL) and sgRNA 5-(40 ng/uL) was prepared and injected (1 nL total volume) into one-cell stage embryos (Ekwill strain). Cutting efficiency was assessed by pooling injected embryos at 24 hr post fertilization (hpf), extracting genomic DNA, amplifying the genomic targeted region by PCR (see Supplementary file 1B for primers), and using the Cel1 assay (Oleykowski et al., 1998) to detect evidence of targeted mutations.

To pinpoint the exon at which skeletal muscle disarray occurs, recombinant Cas9 and high dose gRNAs (100 ng/uL) corresponding to exons 113, 114, 115, 116, 122 and the predicted start codon for the alternative promoter were each injected into ∼120 one-cell stage embryos. Given that homozygotes of either N- or C-terminal truncation mutants of ttna cause cessation of cardiac contraction, we focused our attention on those fish with obvious contractile defects and further phenotyped these through motility assays and immunohistochemistry (see below). For each of the targeted exons, 10 embryos with cessation of contraction were analyzed by immunostaining.

Husbandry and genotyping

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Zebrafish founders were outcrossed onto the Ekwill background, and subsequent progeny outcrossed again. F2 founders were then in-crossed to obtain F3 generation homozygote mutants, which were used for phenotypic and genomic analysis. Genotyping was performed using the Cel1 assay (Oleykowski et al., 1998) and/or allele-specific PCR analysis (Gaudet et al., 2007).

Immunohistochemistry

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Embryos were fixed and stained using standard procedures (Deo et al., 2014). Briefly, anesthetized embryos at 3 dpf were fixed in 4% paraformaldehyde (PFA) for 12 hr. Fixed tissue was washed in PBS + 0.1% Triton X-100 (PBST), permeabilized with 10 ug/mL proteinase K in PBST, and then refixed with 4% PFA for 20 min after which it was subjected to standard blocking and antibody incubations. Primary antibodies used were mouse anti-human alpha-actinin (Life Technologies, MA1-22863, 1:500 dilution) and titin T11 (Sigma-Aldrich, St. Louis, MO, T9030, 1:100) and T12 (1:10, a generous gift from Elisabeth Ehler and Dieter Fuerst). The secondary antibody was donkey-anti-mouse conjugated to an Alexa-488 fluorophore (Life Technologies, R37114, 1:500).

Imaging and data analysis

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Videos at 100–250 frame per second of beating hearts were taken from live 72 hr post-fertilization (hpf) embryos using an Axiozoom V16 Stereo (Carl Zeiss, Oberkochen, Germany) with a 10× objective lens and an Orca Flash 4.0T high-speed digital camera (Hamamatsu Photonics, Hamamatsu City, Shizuoka, Japan). Confocal images were collected on a Leica TCS SP5 with a 63x objective lens. To assess motility, embryos were tapped gently with a whisker.

Morpholino knockdown

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ATG- and splice-blocking morpholinos to ttna (conventional and alternative promoter) and ttnb were designed with the help of Gene Tools staff (http://www.gene-tools.com/). Morpholino sequences were TAA ATG TTG GAG CTT GCG TTG ACA T (ATG) and AAT TAC CTT TGA TTG TGA CTT TGG T (Seeley et al., 2007) for ttnb and CCT GTG TTG TCA TGG TGG AAG GCA C (conventional ATG), GTT CCA GGA GAC ACA GGT AAT CCC T (internal ATG) and TGC AAC TGA TAC TCA CCT TCT CCA C (exon4-intron4 junction) for ttna. Morpholinos were re-suspended in sterile water to a concentration of 1 mM and diluted to working concentration in sterilized water. Male and female wild-type adult zebrafish were housed and embryos bred using standard protocols (Westerfield, 1993). Morpholinos were introduced into the zebrafish yolk via microinjection at no later than the two-cell developmental stage. Injected embryos were then kept at 28.5°C in E3 solution (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4). The conventional promoter ttna/ttnb ATG-morpholinos were highly toxic, even at low concentrations. We thus focused on using the ttna and ttnb splice-blocking morpholino’s as well as the internal promoter ttna ATG morpholino at working concentrations of 5 ng/nl. To further reduce toxicity, we co-injected the ttna splice-blocking morpholino with a previously described p53 morpholino at 5 ng/nl (Robu et al., 2007). In terms of sample size, we typically injected 100–150 embryos for every experiment.

Zebrafish RNA-sequencing and analysis

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High depth RNA-Sequencing of wild-type embryonic hearts was first performed to select constitutive exons for gRNA targeting. Embryo heart purification was carried out as described (Geoffrey Burns and Macrae, 2006). Briefly, 300–400 embryos of Tg (cmcl2-GFP) at 3 dpf were pooled for a single sequencing experiment. Embryo Dissociation Medium (EDM) was freshly prepared using Leibovitz’s L-15 Medium (Life Technologies, 11415–064) containing 10% FBS (Sigma, F-2442), and kept ice-cold prior to use. After washes with EDM, embryos in 1 ml of EDM were drawn to a 3 ml syringe with 19G needle and expelled back to the tube for 30 times. Large embryo fragments were filtered out using a 100 um filter, and the filter was washed several times with more EDM. The flow-through was collected in a 60-mm glass petri dish on ice, and then applied to a 40 um filter. The filter was inverted and the retained fragments including hearts were washed off into a glass dish with 5x1ml EDM. GFP + hearts were sorted under fluorescent light using a Leica (Leica Camera, Wetzar, Germany) M205 FA fluorescence microscope. Trizol-extracted RNA was purified by column (RNA Clean and Concentrator, Zymo Research, R1016). Sequencing libraries were prepared using the SureSelect Strand Specific RNA-Seq kit (Agilent Technologies, Santa Clara, CA, G9691A), following the manufacturer’s instructions. 101 bp paired end sequencing was performed on a HiSeq 2500 sequencer (Illumina, San Diego, CA).

For analysis of splicing of ttna mutants, hearts and skeletal muscle (trunk) were isolated from ttna mutant and wild-type zebrafish embryos (72 hrpf) and adults using manual dissection after euthanasia. Pooled hearts and skeletal muscle from 10 fish were used for subsequent library preparation. The tissue was lysed in Trizol (Life Technologies). RNA was recovered from the aqueous phase and purified purified by column (RNA Clean and Concentrator, Zymo Research, R1016) according to manufacturer’s protocol. cDNA was prepared using the Ovation RNA-Seq System V2 (NuGEN Technologies) with sonication into 200bp fragments using an M220 sonicator (Covaris, Woburn, MA). Sequencing libraries were prepared using the Ovation Ultralow Ultralow DR Library System (NuGEN Technologies, San Carlos, CA). 101 bp single end sequencing was performed on a HiSeq 2500 sequencer in rapid run mode, with 7 samples per lane of the flow cell.

Reads were mapped to the dr7 build of the zebrafish genome using TopHat (Trapnell et al., 2009). Samples were normalized using the edgeR package (Robinson et al., 2010), with trimmed mean of M values (TMM) normalization. Percent spliced in estimates were made using in-house Python scripts implementing previously published methods of splice isoform quantification using exon-exon junction reads (Pervouchine et al., 2013), which avoid the estimation difficulties arising from non-uniform read coverage of exons (Kakaradov et al., 2012).

Mouse RNA-sequencing and analysis

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The heart from an E12.5 129X1/SvJ x C57BL/6 cross mouse was disrupted in Trizol with plastic pestle and passed through QIAshredder (Qiagen, Hilden, Germany, 79654) for homogenization. RNA was isolated from Trizol using a 1:1 mixture with 70% ETOH followed by column purification and DNase treatment (RNeasy Mini kit, Qiagen, 74104). The cDNA was generated and amplified using the Ovation RNA-Seq System V2 (NuGEN Technologies) and the library was prepared using the Ovation Ultralow Ultralow DR Library System (NuGEN Technologies). 101 bp single end sequencing was performed on a HiSeq 2500 sequencer. Reads were mapped to the mm9 build of the mouse genome using TopHat (Trapnell et al., 2009).

Nonsense-mediated decay rate estimates

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To estimate nonsense-mediated decay (NMD) rates, we evaluated allelic imbalance in heterozygote embryos. For each mutant of interest, total RNA was extracted from three heterozygote embryos. Primers, designed to span at least one exon-exon junction, were used to amplify ∼100 bp surrounding the mutation of interest. For each embryo, 2 independent PCR reactions - each with 10 amplification cycles - was performed. Next generation sequencing libraries were prepared using the NEBNext Ultra kit (New England Biolabs, Ipswich, MA, E7370S) with 10 cycles post-amplification. Samples were assessed by 100 bp single end sequencing on a MiSeq instrument (Illumina). The ratio of mutant to wild-type reads was used as a measure of allelic imbalance. For each mutation, between 600 and 10,000 reads were used for ratio estimates.

Quantitative real-time PCR to assess ratio of Cronos to full-length ttna/Ttn in zebrafish and mouse

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Forward primers mapping to either e115 or the Cronos isoform was used in combination with a reverse primer spanning the e116-e117 exon-exon junction. Similar primers were designed to detect Cronos vs. full-length Ttn levels in mouse. Standard curves using a cloned PCR product were used to compare relative primer efficiencies (data not shown). Quantitative real time-PCR analyses were carried out with cDNA templates using SensiFast SYBR green (Bioline Reagents, London, UK) on the CFX384 Real Time PCR system (BioRAD, Hercules, CA) (Heredia et al., 2013).

Zebrafish samples for qPCR consisted of embryonic and adult hearts and trunk muscle, and were isolated as described above. Mouse heart and skeletal muscle samples were taken from C57BL/6 or CD1 mice purchased from Jackson laboratory. All mice were euthanized by CO2 inhalation and cervical dislocation prior to surgical dissection of relevant tissues.

Human RNA-Sequencing analysis

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We downloaded fastq format files for DCM patients and healthy controls corresponding to GEO Accessions GSM1380718, GSM1380719, GSM1380722, and GSM1380723. Reads were mapped to the hg19 build of the human genome using TopHat (Trapnell et al., 2009). PSI values were computed as described above.

Use of Publicly Available DNAse-Seq and ChIP-Seq data

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We downloaded bed format files corresponding to GEO Accessions GSM665817 (fetal heart), GSM530661 (fetal heart), GSM665817 (fetal heart), GSM665824 (fetal heart), GSM665830 (fetal heart), GSM701533 (fetal trunk), GSM772735 (fetal heart), GSM906406 (left ventricle), GSM1027322 (pediatric heart), GSM1058781 (fetal leg), GSM1160200 (fetal trunk), and GSM530654 (fetal heart). Depth of coverage analysis was performed using in-house R scripts and data plotted using the ggplot2 package (Wickham, 2009).

5’ RACE

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5’ RACE was performed using published protocols (Picelli et al., 2014; Matz et al., 1999). An oligonucleotide for template switching (TSO, Univ_UMI_RACE, Supplementary file 1B) was purchased from Exiqon (Woburn, MA). Total RNA was obtained from three sources. Zebrafish RNA was isolated from zebrafish skeletal muscle, as described above. Mouse RNA was isolated from cardiac tissue of a euthanized adult mouse (C57BL/6) using manual dissection of the heart followed by tissue homogenization, Trizol extraction, and RNA column purification (RNA Clean and Concentrator, Zymo Research, R1016). Fetal human heart (31 week old, male) RNA was purchased (BioChain Institute, Newark, CA, R1244122-50). First strand synthesis was performed using a TTN gene-specific primer (Supplementary file 1B) and the TSO, with SmartScribe reverse transcriptase (Clontech, Mountain View, CA, 639536) and with added SUPERase In RNAse inhibitor (Life Technologies, AM2694). Uracil-DNA Glycosylase (New England Biolabs, M0280S) was used to degrade any remaining TSO according to manufacturer’s instructions. PCR amplification of the resulting cDNA was performed using a nested gene-specific primer (Supplementary file 1B) and a universal 5’ primer (Supplementary file 1B) with a touch-down annealing temperature protocol. PCR products were gel purified (Zymo Research, D4001), blunt-end cloned using the pGEM-T Easy Vector system (Promega, Madison, WI, A12360) and analyzed by Sanger sequencing.

In situ hybridization

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Whole-mount in situ hybridization with digoxigenin-labeled mRNA antisense probes to Cronos and ttna was performed as previously described (Thisse and Thisse, 2008). For Cronos, an 89nt digoxigenin-labeled riboprobe unique to this isoform was hybridized at 60°C. For ttna, a 453nt riboprobe corresponding to a constitutive region was hybridized at 68°C. In both cases embryos were developed in BM purple (11442074001, Sigma-Aldrich).

Protein electrophoresis

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Briefly, euthanized zebrafish embryos were Dounce homogenized in lysis buffer (500 mM Tris pH 7.4 150 mmM NaCl, 1% NP-40, 1 mM PMSF, 1x protease inhibitor cocktail [Sigma-Aldrich]) on ice and centrifuged at 14,000 g for 10 min at 4C. The denaturing/loading buffer (8 M urea, 2 M thiourea, 3% SDS, 75 mM DTT, 0.1% bromophenol blue, 0.05% Tris-HCl pH 6.8) was mixed with the lysate at a 4:1 ratio and the resulting sample denatured at 60°C for 15 min. Electrophoresis was performed using a denaturing 1% agarose gel at 15mA for 3–6 hr (3 hr for Myosin and 6 hr for Titin) on a Hoefer (Holliston, MA) SE 600 gel unit (Steffen, et al., 2007Warren, et al., 2003). Gels were fixed for at least 1 hr after running, briefly washed in water and vacuum dried at 40C overnight. The following day, the gel was rehydrated and stained with SYPRO Ruby protein gel stain (S4942, Sigma-Aldrich) according to manufacturer’s instructions.

Senior athlete recruitment

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199 healthy senior athletes with no prior history of cardiac disease were recruited between 2004 and 2012 at the Huntsman World Senior Games (held in St George, Utah). DNA from whole blood was extracted using the Gentra Puregene DNA isolation kit (Qiagen, 158445).

Targeted capture-based sequencing and variant calling

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Targeted capture and variant calling were performed as previously described (Deo, et al., 2014). Briefly, NimbleGen’s SeqCap EZ Choice technology was used to construct oligonucleotide probes complementary to the exons (Ensembl GRCh37) of 160 genes involved or predicted to be involved in Mendelian forms of cardiac disease (Supplementary file 1A). Genomic DNA libraries were sheared by sonication on a M220 sonicator (Covaris), hybrdized to the capture probes, and libraries prepared from the captured material using the Kapa Biosystems (Wilmington, MA) Library Preparation Kit. Libraries were sequenced on an Illumina HiSeq 2500 sequencer with multiplexing of 24 samples per lane. Assembly was performed using bwa (Li and Durbin, 2009) and variants called using the GATK (McKenna, et al., 2010). Median coverage was 455-fold with 99.6% of exons found to be callable for the major cardiac N2BA isoform (ENST00000591111)and 98.4% callable for the Inferred Complete isoform (ENST00000589042). Seven samples were captured with a prior cardiomyopathy gene panel of 116 genes (Deo, et al., 2014).

Coding sequence mutations resulting in a premature nonsense codon or frame-shift were considered to be truncating mutations. For mutations of canonical (-1/-2) splice acceptor variants, which would most likely result in exon skipping, we further required that the downstream exon was not in the same codon phase as the skipped exon, as this would fail to produce a frame-shift and thus not result in a truncation (such a situation is common in the I-band). Likewise, for mutations of canonical splice donor variants (+1/+2), which we predicted would result in intron read-through, we performed in silico translation of the resulting retained intron to ensure that a premature truncation was observed (this was not always the case, especially for the I-band).

Statistical analysis

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All statistical analysis was performed in R (3.1.1). A difference in proportion test was applied to the prevalence of TTN truncations in senior athletes vs. literature controls, as well as the fraction of TTN truncations that map to the Novex-3 exon.

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Decision letter

  1. Harry C Dietz
    Reviewing Editor; Howard Hughes Medical Institute and Institute of Genetic Medicine, Johns Hopkins University School of Medicine, United States

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled "An internal promoter underlies the difference in disease severity between N- and C-terminal truncations of Titin in zebrafish" for peer review at eLife. Your submission has been evaluated by Mark McCarthy (Senior Editor) and three reviewers, one of whom, Harry Dietz, is a member of our Board of Reviewing Editors. Your manuscript has generated considerable enthusiasm. The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

This interesting manuscript provides a provocative explanation for a well-described phenotype-genotype correlation in dilated cardiomyopathy (DCM) caused by mutations in the gene encoding titin (TTN). Specifically, C-terminal heterozygous truncating mutations associate with a much more severe presentation of DCM than do N-terminal truncating mutations, with an apparent discriminating boundary between exons 115 and 116. In this manuscript, the authors recapitulate this experience using a series of targeted mutations in the ttna gene in zebrafish, which also have a paralogous ttnb gene. Evidence is provided to suggest that these observations are not based upon alternative splicing, differential efficiency of nonsense-mediated mRNA decay (NMD) or differential dominant-negative potential of the predicted truncated protein products derived from the mutant alleles. Instead, the data strongly suggest the relevance of an alternative promoter that lies between exons 115 and 116, giving rise to an mRNA and protein (termed Cronos) that is expressed (and putatively functional) with proximal but not distal premature termination codon mutations. Overall, this is a very elegant study with quality data and largely convincing conclusions. However, there are a number of issues and questions that need to be addressed before we can consider this further.

1) The most substantive concern expressed by multiple reviewers relates to the relative strengths and limitations of the zebrafish model. The lack of correlation between location of truncating mutations and severity of cardiomyopathy in this model is inadequately explored and discussed. A comprehensive tissue-specific quantitative and developmental survey of alternative promoter utilization in both heart and skeletal muscle is indicated. The rationale for studying homozygous variation in zebrafish is somewhat difficult to justify completely and has an impact on the generalizability of the results. The authors should consider higher resolution phenotyping of heterozygous animals with or without environmental stressors. They should also comprehensively describe (and perhaps illustrate) what is known regarding species-specific TTN genomic structure and message biology.

2) NMD is generally a very efficient process. There is generally an apparent polarization effect, with greater efficiency for proximal PTCs, with the potential for less efficiency for distal PTCs and near-universal loss of NMD for PTCs in the very distal portion of the penultimate exon or in the last exon. In this light, it is surprising that all assayed PTCs (none in last exons) had a relatively modest effect on transcript abundance (∼50% of WT). The details regarding how this assay was performed are minimal and should be expanded. Would the assay detect the Cronos transcript that is expected to be present for N- but not C-terminal truncations? If so, why is this not reflected in the results of this experiment?

3) Additional experimental details and/or controls are also needed for the dominant-negative assay, for example the control for efficient morpholino-induced exon skipping related to observation of cardiomyopathy in WT mice. There needs to be some demonstration of high efficiency in skeletal muscle. Also, the authors should consider the possibility of a dosage-dependent dominant-negative effect when interpreting their results.

4) The apparent demonstration of the Cronos protein product (Figure 5) is not convincing. There is poor resolution, no demonstration of specificity, no identification of full-length protein, no explanation for the variation in intensity of other protein isoforms that vary between genotypes, and no loading control.

5) Much more detail is needed regarding the exact location and predicted functional consequence of the truncating mutations observed in senior athletes vs. various control populations. When compared to healthy volunteers or participants in heart study cohorts (numbers not specified), senior athletes had about the same frequency (∼1.0-1.5%) of TTN truncation mutations. However, all senior athletes had their truncating mutation in the very distal portion of the Novex-3 isoform (that is rather poorly described in this manuscript), while the other groups largely showed ‘other’ types of truncations. Apparently, there is an alternative TTN transcript that includes a large (∼6.5kb) terminal exon that also contains a 3' cleavage and polyadenylation signal. By report, the Novex-3 transcript contains all proximal exons, and terminates considerably short of the alternative promoter described here. Are the truncations in the very distal portion of the Novex-3 exon seen in robust old age because they have no functional impact at all (i.e. normal levels of both full-length and Novex-3 message and protein due to avoidance of NMD in both)? While the data might even suggest a selective advantage, there is no mechanism that can even be inferred by the data and conclusions of this manuscript. Perhaps more importantly, what was the specific nature of the ‘other’ truncating mutations in the healthy volunteers and other unselected cohorts? Were they greatly enriched for N- as opposed to C-terminal truncations (defined by the exon 115/116 boundary), as might be predicted by this study? Were the Novex-3 truncations seen in the FHS and JHS cohorts also in the extreme C-terminal exon, or were they more widely distributed?

6) There are concerns regarding generalizations made about existing TTN phenotype-genotype correlations in human cardiomyopathy patients. The initial inference that C-terminal titin truncation variants are uniformly classified as pathogenic mutations is incorrect. There is a substantial excess of C-terminal truncating variants in DCM subjects, but there are also such variants in the control groups so the position of a variant is insufficient (at least at this resolution) to define pathogenicity. This observation also raises important questions regarding the mechanisms of pathogenicity. In addition, it is worth noting that there are remarkably few of the proposed truncating mutations that have been demonstrated to result in truncated protein and there are only two extant families where there are any segregation data of significance between titin variants of any sort and the DCM phenotype. In each case, the variant is not clearly truncating when studied closely. Finally, the spectrum of variation in titin that leads to skeletal muscle involvement is not so systematically described but many DCM subjects (with heterozygous genotypes) have subclinical skeletal muscle involvement when examined rigorously. The suggestion that TTN mutations cause HCM is controversial and should be stated as such. The distinction between N- and C- terminal truncating mutations with regard to DCM is not as discrete and robust as is suggested by the authors, with documentation of cases with N-terminal truncations (i.e. Herman DS et al, NEJM 2012). Also, given the complexities of TTN transcript biology that remain to be fully explored in fish, mice and especially people, the authors’ suggestion that it is now possible to definitively classify C-terminal variation in the titin gene is overstated. While this work will undoubtedly contribute to progress, much more needs to be done to reliably inform patient management and counseling.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "An internal promoter underlies the difference in disease severity between N- and C-terminal truncations of Titin in zebrafish" for further consideration at eLife. Your revised article has been favorably evaluated by Mark McCarthy (Senior Editor), Harry Dietz (Reviewing Editor), and two reviewers. The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance, as outlined below:

Please take note of the additional comments by reviewer #3 and specifically further highlight the complexity and therefore inherent limitations of the zebrafish system.

Reviewer #2:

This revised manuscript addresses all of my prior comments well. The manuscript is substantially improved. I have no additional concerns or recommendations.

Reviewer #3:

The authors suggest that they have clarified the relationship between zebrafish and human titin biology, but they have simply fitted their observations to a model where homozygous titin null zebrafish skeletal muscle is representative of human heterozygous null cardiac muscle. While this may be the case, the fact is that sarcomerogenesis itself is disrupted during development in the fish limb of this construct, while humans with the proposed orthologous trait have no demonstrable structural defect in the sarcomere and survive to middle age.

The authors' also assert that components of the observed disparities between fish and human hearts and fish and human skeletal muscle are a consequence of the associated levels of ttnb and Cronos in the different tissues in each species. It would be important to formally demonstrate these assertions with rescue experiments in the different tissues, or as noted in prior reviews, to consider and test alternative explanations including the presence or absence of external stressors. The authors did test two rather acute and rather similar perturbations without success, though the zebrafish would seem to be amenable to much more extensive and stringent challenge.

The RNAseq data more robustly quantitate the levels of NMD and are reassuring, though the distribution along the transcript is somewhat discordant with prior observations in other genes.

I remain somewhat unsure how to interpret the failure of ttna knockdown to rescue the ttna homozygous mutants. It is difficult to see how a mechanism that may be dependent on the presence of a threshold amount of wild-type (WT) protein, can be excluded by the knockdown of both mutant alleles. This serves only to emphasize the failure of the zebrafish experiments as described to capture the biology of either human skeletal or cardiac titin disease.

The authors have revised their statements on the inferences of their work for clinicians and in doing so have created an improved and carefully nuanced description of the field.

https://doi.org/10.7554/eLife.09406.024

Author response

This interesting manuscript provides a provocative explanation for a well-described phenotype-genotype correlation in dilated cardiomyopathy (DCM) caused by mutations in the gene encoding titin (TTN). Specifically, C-terminal heterozygous truncating mutations associate with a much more severe presentation of DCM than do N-terminal truncating mutations, with an apparent discriminating boundary between exons 115 and 116. In this manuscript, the authors recapitulate this experience using a series of targeted mutations in the ttna gene in zebrafish, which also have a paralogous ttnb gene. Evidence is provided to suggest that these observations are not based upon alternative splicing, differential efficiency of nonsense-mediated mRNA decay (NMD) or differential dominant-negative potential of the predicted truncated protein products derived from the mutant alleles. Instead, the data strongly suggest the relevance of an alternative promoter that lies between exons 115 and 116, giving rise to an mRNA and protein (termed Cronos) that is expressed (and putatively functional) with proximal but not distal premature termination codon mutations. Overall, this is a very elegant study with quality data and largely convincing conclusions. However, there are a number of issues and questions that need to be addressed before we can consider this further. 1) The most substantive concern expressed by multiple reviewers relates to the relative strengths and limitations of the zebrafish model. The lack of correlation between location of truncating mutations and severity of cardiomyopathy in this model is inadequately explored and discussed. A comprehensive tissue-specific quantitative and developmental survey of alternative promoter utilization in both heart and skeletal muscle is indicated. The rationale for studying homozygous variation in zebrafish is somewhat difficult to justify completely and has an impact on the generalizability of the results. The authors should consider higher resolution phenotyping of heterozygous animals with or without environmental stressors. They should also comprehensively describe (and perhaps illustrate) what is known regarding species-specific TTN genomic structure and message biology.

With regard to the primary area of contention, we recognize that we have done an inadequate job drawing a path between our observations in the zebrafish model and human cardiac disease. In the revised manuscript we have sought to make a clearer, more compelling case with the following changes:

A) Comparison of zebrafish and human titin:

The following text is included verbatim in the revised manuscript and we have included a new Figure (Figure 1–figure supplement 1) to graphically illustrate the domain organization of human TTN and zebrafish ttna and ttnb, as suggested. Our primary objectives in this section are to 1) highlight the similarities between the human and zebrafish Titin proteins, and 2) emphasize that the paralogous ttna and ttnb proteins are both essential for skeletal muscle sarcomere development, creating a scenario where, at least in skeletal muscle, studying a homozygote knockout of one of these genes bears resemblance to studying a heterozygote in mammals.

We have also rephrased the text accordingly. Please see: “As a result of an ancestral genome duplication event, zebrafish include two titin genes: ttna and ttnb […] severe disruption of skeletal muscle sarcomeric architecture, highlighting the mutual contribution of the two genes (14).”

B) Tissue-specific quantitative and developmental survey of Cronos expression:

In the revised manuscript, we have added 3 new figure panels that characterize Cronos expression in zebrafish and mouse by whole mount in situ hybridization (with one probe specific to Cronos) and by quantitative real-time PCR (Figure 6A-C). Our primary observation is that Cronos expression is at high levels in zebrafish skeletal muscle during development (∼2:1 ratio of Cronos to full-length ttna), but drops off dramatically during adulthood (∼1:70 ratio). Although the Cronos expression in zebrafish heart also appears to be developmentally regulated, it is much lower than in skeletal muscle, with a ∼1:5 ratio at 72 hr and 1:30 ratio in adulthood.

In contrast, the relative levels of Cronos in mouse heart are much higher than in skeletal muscle, with nearly a 1:1 ratio of Cronos to full-length Titin in E12.5 heart. Thereafter mouse cardiac expression of Cronos decreases to ∼20–30% of full-length Titin. Skeletal muscle expression varies widely across development and in different muscle beds with a 1:5 ratio in mouse hindlimb at P2, and between 1:16 and ∼1:1000 ratios in adult extensor digitorum longus (fast-twitch) and soleus (slow-twitch) muscle, respectively.

We indicate that the poor expression of Cronos in zebrafish heart can be a possible explanation for the indistinguishable cardiac phenotypes of N- and C-terminal truncation mutants, though the fact that ttnb plays a minimal role in cardiac development is probably also contributing to the severity of the ttna mutants.

In addition to the additional Figure 6, we have included two paragraphs summarizing the tissue-specific and developmental stage results: “We next surveyed the expression levels of Cronos in cardiac and skeletal muscle […] expression varies across development and in different muscle beds with a 1:5 ratio in mouse hindlimb at P2, and between 1:16 and ∼1:1000 ratios in adult extensor digitorum longus (fast-twitch) and soleus (slow-twitch) muscle, respectively.”

C) Interpretation of lack of variation in truncation mutation severity in heart:

We attribute the stark differences between skeletal muscle phenotypes in ttna N- and C-terminal truncation mutants to 1) the prominent expression of Cronos in developing skeletal muscle and 2) the presence of an important functional paralog, ttnb, which can substitute for ttna function. Unfortunately, in cardiac muscle the contribution of both of these factors is severely diminished, including low expression of Cronos, and a more minimal role for ttnb in cardiac sarcomerogenesis.

We are reassured that there is prominent expression of Cronos in developing mouse heart, which is in keeping with Cronos playing an important role in mammalian heart development and therefore consistent with the lack of Cronos intensifying DCM phenotypic severity. Interestingly, others have described a prominent band on protein gels of developing mouse and rat heart tissue at the expected molecular weight of Cronos, which is substantially diminished in adulthood. This band has been dismissed as a degradation product – but we speculate that in some cases this is very likely to be Cronos.

We touch on the limited cardiac expression of Cronos in zebrafish as well as the limited contribution of ttnb in heart development as explanation for our zebrafish cardiac results. We also include an additional paragraph in the Discussion: “Our work also proposes an alternative explanation for the consistently observed ‘T2’ band on protein gels, which migrates at the expected molecular weight of Cronos. This has invariably been labeled a C-terminal degradation product, as it reacts with A-band antibodies (25). However, in keeping with Cronos expression patterns in development, the T2 band was prominently seen in developing mouse (25) and rat hearts (26), but was markedly reduced in adulthood. It is possible that, at least in some situations, T2 and Cronos represent the same isoform.”

D) Perturbation induced discrimination of N- and C-terminal heterozygotes:

We agree that perturbation experiments in heterozygotes (such as was done with Ttn+/- mice, Gramlich et al, 2009) would have been more consistent with the human disease scenario, although the presence of paralogs complicates this situation in zebrafish. Although we tried isoproterenol (and angiotensin II) at several doses and at several developmental time points and used natriuretic peptide levels as readout, we were not able to consistently distinguish WT fish from heterozygotes (N- or C). Given the low cardiac expression of Cronos, we suspect that even with an optimized assay, the distinction of N- and C-terminal truncation mutants from one another would be very challenging. 2) NMD is generally a very efficient process. There is generally an apparent polarization effect, with greater efficiency for proximal PTCs, with the potential for less efficiency for distal PTCs and near-universal loss of NMD for PTCs in the very distal portion of the penultimate exon or in the last exon. In this light, it is surprising that all assayed PTCs (none in last exons) had a relatively modest effect on transcript abundance (∼50% of WT). The details regarding how this assay was performed are minimal and should be expanded. Would the assay detect the Cronos transcript that is expected to be present for N- but not C-terminal truncations? If so, why is this not reflected in the results of this experiment?

This is an excellent point. Our prior approach to NMD estimate was inadequate and involved normalizing total ttna to ttnb levels in homozygote ttna null mutants. In the revised manuscript we have performed targeted RNA-Seq of the mutant exons in heterozygote embryos to estimate NMD by looking at allelic imbalance (Figure 2D). Comparing raw counts of mutant and WT alleles, we see ∼80% of the mutant transcript degraded, in keeping with established NMD efficacy. Interestingly, we see very little variation across the length of the transcript. We have also performed qPCR to estimate the ratio of Cronos to full-length ttna in N-terminal and C-terminal mutants and find the expected result (Figure 5B): a high ratio of Cronos to full-length ttna for all N-terminal mutants and a much lower rate, more comparable to WT, for the C-terminal mutants. We suspect that some variation in promoter activity for the full-length and Cronos transcripts could account for some of the variation in these ratios within the N- and C-terminal mutant groups.

We have modified the text and figures to include this data, which we believe strengthens our story. For example, we now state: “Given that NMD substantially reduces the levels of ttna transcripts with premature termination codons (Figure 2D), we reasoned that we should be able to distinguish N- and C-terminal truncation mutants by the ratio of Cronos to full-length transcript. In ttnan/n mutants, which would degrade full-length ttna but not Cronos, we would anticipate a higher Cronos:full-length ratio than in ttnac/cmutants mutants, which should degrade both transcripts approximately equally. As expected, we see a much higher ratio of Cronos:full-length transcript levels in ttnan/nmutants than in ttnac/cmutants (Figure 5B).” 3) Additional experimental details and/or controls are also needed for the dominant-negative assay, for example the control for efficient morpholino-induced exon skipping related to observation of cardiomyopathy in WT mice. There needs to be some demonstration of high efficiency in skeletal muscle. Also, the authors should consider the possibility of a dosage-dependent dominant-negative effect when interpreting their results.

We have now performed qPCR to demonstrate ∼78% knockdown efficiency using our ttna morpholino. Either exon skipping or intron retention arising from morpholino injection should result in an N-terminal truncation on top of the C-terminal mutation. In conjunction with NMD, this intervention should result in residual C-terminal truncated protein being at ∼5% of the level of ttnb. Given that we see no amelioration in skeletal muscle function (fish remain immobile) or architecture, it is difficult to reconcile this observation with a dominant negative mechanism. Similarly, for dosage to be responsible for the difference between N- and C-terminal truncations, we would not expect to have seen similar levels of NMD for N- and C-terminal truncations nor see persistence of skeletal muscle pathology with such a low level of mutant C-terminal protein present.

In the manuscript, we have made the following changes: “Knockdown by morpholino was effective (∼78% knockdown efficiency, data not shown) and the combined action of NMD and morpholino knockdown would be expected to reduce mutant ttna protein to only ∼5% of the level of ttnb protein. Nonetheless we were unable to rescue skeletal muscle architecture in ttnac/cmutants (Figure 3B). We thus concluded that ttnac/c mutants do not act as dominant negatives and that an alternative explanation was needed.”

4) The apparent demonstration of the Cronos protein product (Figure 5) is not convincing. There is poor resolution, no demonstration of specificity, no identification of full-length protein, no explanation for the variation in intensity of other protein isoforms that vary between genotypes, and no loading control.

We have updated this gel to include morpholino knockdown controls as well as loading controls. We have found this gel technically challenging to execute, potentially because of the long running time ∼6 required, but we believe the results are now convincing.

5) Much more detail is needed regarding the exact location and predicted functional consequence of the truncating mutations observed in senior athletes vs. various control populations. When compared to healthy volunteers or participants in heart study cohorts (numbers not specified), senior athletes had about the same frequency (∼1.0–1.5%) of TTN truncation mutations. However, all senior athletes had their truncating mutation in the very distal portion of the Novex-3 isoform (that is rather poorly described in this manuscript), while the other groups largely showed ‘other’ types of truncations. Apparently, there is an alternative TTN transcript that includes a large (∼6.5 kb) terminal exon that also contains a 3' cleavage and polyadenylation signal. By report, the Novex-3 transcript contains all proximal exons, and terminates considerably short of the alternative promoter described here. Are the truncations in the very distal portion of the Novex-3 exon seen in robust old age because they have no functional impact at all (i.e. normal levels of both full-length and Novex-3 message and protein due to avoidance of NMD in both)? While the data might even suggest a selective advantage, there is no mechanism that can even be inferred by the data and conclusions of this manuscript. Perhaps more importantly, what was the specific nature of the ‘other’ truncating mutations in the healthy volunteers and other unselected cohorts? Were they greatly enriched for N- as opposed to C-terminal truncations (defined by the exon 115/116 boundary), as might be predicted by this study? Were the Novex-3 truncations seen in the FHS and JHS cohorts also in the extreme C-terminal exon, or were they more widely distributed?

We have modified this section and included 2 new figure panels: 1) one that depicts the position of mutations in DCM patients and control populations or from the senior athletes onto a schematic of the TTN meta-transcript 2) a ratio of mutations found in regions C-terminal to the Cronos promoter to those in the N-terminus in different populations. We have eliminated the figure panel that emphasizes the location of senior athlete mutations to the terminal exon of the Novex-3 isoform. In fact, when we referred to the Novex-3 isoform we meant only those mutations mapping to the terminal exon, which is unique to this isoform.

The following text is now included in the revised manuscript: “We compared the frequency and distribution of TTN truncation mutations to those found in unselected controls and healthy volunteers from a recent sequencing study (CTLlit) (7) […] As expected, in end-stage DCM we observed a much higher rate of mutants C-terminal to Cronos than N-terminal (30:1), followed by unselected DCM (37:11), literature controls (6:11, 2:2, 10:7 for FHS, healthy volunteers and JHS) and senior athletes (0:3).” 6) There are concerns regarding generalizations made about existing TTN phenotype-genotype correlations in human cardiomyopathy patients. The initial inference that C-terminal titin truncation variants are uniformly classified as pathogenic mutations is incorrect. There is a substantial excess of C-terminal truncating variants in DCM subjects, but there are also such variants in the control groups so the position of a variant is insufficient (at least at this resolution) to define pathogenicity. This observation also raises important questions regarding the mechanisms of pathogenicity. In addition, it is worth noting that there are remarkably few of the proposed truncating mutations that have been demonstrated to result in truncated protein and there are only two extant families where there are any segregation data of significance between titin variants of any sort and the DCM phenotype. In each case, the variant is not clearly truncating when studied closely. Finally, the spectrum of variation in titin that leads to skeletal muscle involvement is not so systematically described but many DCM subjects (with heterozygous genotypes) have subclinical skeletal muscle involvement when examined rigorously. The suggestion that TTN mutations cause HCM is controversial and should be stated as such. The distinction between N- and C- terminal truncating mutations with regard to DCM is not as discrete and robust as is suggested by the authors, with documentation of cases with N-terminal trucations (i.e. Herman DS et al, NEJM 2012). Also, given the complexities of TTN transcript biology that remain to be fully explored in fish, mice and especially people, the authors' suggestion that it is now possible to definitively classify C-terminal variation in the titin gene is overstated. While this work will undoubtedly contribute to progress, much more needs to be done to reliably inform patient management and counseling.

We apologize for creating the impression that our work has resulted in newfound certainty in genotype-phenotype correlations. From our experience taking care of many dozens of families with inherited cardiomyopathies, such determinism is never the case. We have moderated language that makes any such suggestion in the revision. We would like to address some of the above comments and concerns:

1) The presence of ‘controls’ with C-terminal truncations does not eliminate the possibility that these mutations do in fact have cardiac consequences that fall short of overt heart failure, which was the motivation for studying a hyper-normal senior athlete population. Finding C-terminal truncations in the senior athlete population would have cast greater doubt on whether such mutations are sufficient to compromise cardiovascular fitness.

2) We are encouraged by this report of subclinical skeletal manifestations in DCM TTN patients though we cannot find any published studies that have rigorously documented this.

3) We have eliminated mention of the HCM association with TTN.

4) Our work actually does not address if N-terminal truncations – especially in constitutive exons – are deleterious. We analyzed the senior athletes with the hope that we might find N-terminal truncations though small sample size precludes robust conclusions. We do note that in Supplementary Table 12 of Herman et al, the dozen or so families with evidence of segregation in 2 or more members all had C-terminal truncations.

Please see the following passage in the manuscript: “Our results have two clinical implications for titinopathy patients […] although it is unclear whether there is a restricted developmental window where this might be possible.”

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "An internal promoter underlies the difference in disease severity between N- and C-terminal truncations of Titin" for further consideration at eLife. Your revised article has been favorably evaluated by Mark McCarthy (Senior Editor), Harry Dietz (Reviewing Editor), and two reviewers. The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance, as outlined below: Please take note of the additional comments by reviewer #3 and specifically further highlight the complexity and therefore inherent limitations of the zebrafish system.Reviewer #2: This revised manuscript addresses all of my prior comments well. The manuscript is substantially improved. I have no additional concerns or recommendations.Reviewer #3: The authors suggest that they have clarified the relationship between zebrafish and human titin biology, but they have simply fitted their observations to a model where homozygous titin null zebrafish skeletal muscle is representative of human heterozygous null cardiac muscle. While this may be the case, the fact is that sarcomerogenesis itself is disrupted during development in the fish limb of this construct, while humans with the proposed orthologous trait have no demonstrable structural defect in the sarcomere and survive to middle age. The authors' also assert that components of the observed disparities between fish and human hearts and fish and human skeletal muscle is a consequence of the associated levels of ttnb and Cronos in the different tissues in each species. It would be important to formally demonstrate these assertions with rescue experiments in the different tissues, or as noted in prior reviews, to consider and test alternative explanations including the presence or absence of external stressors. The authors did test two rather acute and rather similar perturbations without success, though the zebrafish would seem to be amenable to much more extensive and stringent challenge. The RNAseq data more robustly quantitate the levels of NMD and are reassuring, though the distribution along the transcript is somewhat discordant with prior observations in other genes. I remain somewhat unsure how to interpret the failure of ttna knockdown to rescue the ttna homozygous mutants. It is difficult to see how a mechanism that may be dependent on the presence of a threshold amount of wild-type (WT) protein, can be excluded by the knockdown of both mutant alleles. This serves only to emphasize the failure of the zebrafish experiments as described to capture the biology of either human skeletal or cardiac titin disease. The authors have revised their statements on the inferences of their work for clinicians and in doing so have created an improved and carefully nuanced description of the field.

In the revised manuscript we have added two paragraphs (shown below) at the beginning of the Discussion to summarize our findings and highlight the strengths and limitations of the system we have used. Although we firmly believe that our findings represent a substantial advance in this field, with strong biological and clinical implications, we recognize the challenges of studying adult onset autosomal dominant diseases with incomplete penetrance in any sort of animal model. Even if one can bring about phenotypic distinctions with artificial perturbations or genetic modifiers or use increasingly sensitive phenotyping methods to capture differences, one cannot be sure such stresses or distinctions are in fact relevant to the human disease process. Furthermore, TTN brings about an even greater challenge because of the technical challenge of performing rescue experiments with a 110 kilobase transgene.

With regard to Reviewer 3’s final comment, we do not understand the statement of how a knockdown of a mutant allele in the setting of a WT alternative (ttnb) is of no value in assessing a dominant negative mode of action and instead “emphasize(s) the failure of the zebrafish experiments as described to capture the biology of either human skeletal or cardiac titin disease”. If one accepts that ttnb is rescuing the ttna null, this is analogous to performing allele specific knockdown of a mutant protein in a heterozygote and looking at phenotypic impact. In the manuscript, we stress: “Loss of function experiments across a number of systems have demonstrated Titin’s importance in diverse aspects of sarcomere development and function, including the ordered assembly of sarcomeric constituents […] Nonetheless, taking all of our results together, we find it a highly plausible explanation that a superimposed deficiency in expression of the Cronos protein product would result in the more severe forms of human cardiac disease seen in DCM patients with C-terminal TTN truncations.”

https://doi.org/10.7554/eLife.09406.025

Article and author information

Author details

  1. Jun Zou

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    JZ, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Diana Tran

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    DT, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  3. Mai Baalbaki

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    MB, Conception and design, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  4. Ling Fung Tang

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    LFT, Conception and design, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  5. Annie Poon

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    AP, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  6. Angelo Pelonero

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    AnP, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  7. Erron W Titus

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    EWT, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6868-9121
  8. Christiana Yuan

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    CY, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  9. Chenxu Shi

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    CS, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  10. Shruthi Patchava

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    SP, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  11. Elizabeth Halper

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    EH, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  12. Jasmine Garg

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    JG, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  13. Irina Movsesyan

    Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    Contribution
    IM, Acquisition of data, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  14. Chaoying Yin

    1. Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, United States
    2. McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, United States
    Contribution
    ChY, Conception and design, Acquisition of data
    Competing interests
    The authors declare that no competing interests exist.
  15. Roland Wu

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Medicine, University of California, San Francisco, San Francisco, United States
    Contribution
    RW, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  16. Lisa D Wilsbacher

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Medicine, University of California, San Francisco, San Francisco, United States
    Present address
    Department of Medicine and Feinberg Cardiovascular Research Institute, Northwestern University Feinberg School of Medicine, United States
    Contribution
    LDW, Acquisition of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  17. Jiandong Liu

    1. Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, United States
    2. McAllister Heart Institute, University of North Carolina at Chapel Hill, Chapel Hill, United States
    Contribution
    JL, Acquisition of data, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  18. Ronald L Hager

    Department of Exercise Sciences, Brigham Young University, Provo, United States
    Contribution
    RLH, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5128-4635
  19. Shaun R Coughlin

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Medicine, University of California, San Francisco, San Francisco, United States
    Contribution
    SRC, Conception and design, Acquisition of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  20. Martin Jinek

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    MJ, Conception and design, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  21. Clive R Pullinger

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Physiological Nursing, University of California, San Francisco, San Francisco, United States
    Contribution
    CRP, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  22. John P Kane

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    JPK, Conception and design, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  23. Daniel O Hart

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, United States
    Contribution
    DOH, Acquisition of data, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  24. Pui-Yan Kwok

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Dermatology, University of California, San Francisco, San Francisco, United States
    3. Institute for Human Genetics, University of California, San Francisco, United States
    Contribution
    PYK, Conception and design, Acquisition of data
    Competing interests
    The authors declare that no competing interests exist.
  25. Rahul C Deo

    1. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, United States
    2. Department of Medicine, University of California, San Francisco, San Francisco, United States
    3. Institute for Human Genetics, University of California, San Francisco, United States
    4. California Institute for Quantitative Biosciences, San Francisco, United States
    Contribution
    RCD, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    rahul.deo@ucsf.edu
    Competing interests
    The authors declare that no competing interests exist.

Funding

American Heart Association (15POST25090054)

  • Jun Zou

National Heart, Lung, and Blood Institute (DP2 HL123228)

  • Rahul C Deo

National Heart, Lung, and Blood Institute (U01 HL107440-03)

  • Rahul C Deo

National Heart, Lung, and Blood Institute (K08 HL093861)

  • Rahul C Deo

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Ethics

Human subjects: Human genetic studies were performed according to institutional guidelines and with the full approval of the University of California San Francisco Committee on Human Research (CHR#10-00207) and all studies performed were in keeping with the original informed consent forms. Informed consent and consent to publish was obtained from all participants.

Animal experimentation: All zebrafish and mouse experimental work conformed to the 'Guide for the Care and Use of Laboratory Animals' published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996). Animal work was performed according to institutional guidelines with the full approval of the University of California Institutional Animal Care and Use Committee (protocols AN090013-03 and AN107039-01).

Reviewing Editor

  1. Harry C Dietz, Howard Hughes Medical Institute and Institute of Genetic Medicine, Johns Hopkins University School of Medicine, United States

Publication history

  1. Received: June 13, 2015
  2. Accepted: October 15, 2015
  3. Accepted Manuscript published: October 16, 2015 (version 1)
  4. Version of Record published: December 1, 2015 (version 2)

Copyright

© 2015, Zou et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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