The Genetics of Heterotaxy Syndrome

Home , C7orf31, GJB6, KIAA0430, KIAA1704, MEOX2, MPP6

A dissertation submitted to the Division of Graduate Studies and Research, University of Cincinnati in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Molecular and Developmental Biology

Jason R. Cowan Master of Science, University of British Columbia, Vancouver, 2007

Committee Chair: Stephanie M. Ware, M.D., Ph.D. Robert B. Hinton Jr., M.D. Linda M. Parysek, Ph.D. S. Steven Potter, Ph.D. Aaron M. Zorn, Ph.D.

Molecular & Developmental Biology Graduate Program College of Medicine, University of Cincinnati Division of Developmental Biology, Cincinnati Children’s Hospital Medical Center Cincinnati, Ohio, USA, 2015 ABSTRACT Congenital heart defects (CHDs) are the greatest cause of infant morbidity and mortality worldwide, occurring in roughly 8 per 1000 live births (~1%). Heterotaxy, a multiple congenital anomaly syndrome resulting from failure to establish left-right (L-R) asymmetry, is characterized by diverse, complex CHDs. Heterogeneous in presentation and etiology, heterotaxy serves as a complex and growing focal point for cardiovascular genetic research.

In the two decades since the zinc finger transcription factor, ZIC3, was first identified as a cause of X-linked heterotaxy, mutations in nearly twenty genes with L-R patterning functions have been detected among patients with heterotaxy. Nevertheless, despite considerable progress, genetic causes for heterotaxy remain largely uncharacterized. With an estimated 70-80% of heterotaxy cases still unexplained, there remains enormous potential for novel gene discovery.

In this dissertation, we have balanced gene discovery efforts aimed at identifying and characterizing novel causes of heterotaxy with studies into the mechanisms governing ZIC3- related heterotaxy. In order to identify novel genetic causes of heterotaxy, array-based single nucleotide polymorphism (SNP) and comparative genomic hybridization (CGH) screens for copy number variation (CNVs) were completed in a large and carefully phenotyped cohort of 225 patients with heterotaxy and CHDs. Identified CNVs with pathogenic potential ranged in size from large unbalanced translocations to smaller, kilobase-scale abnormalities. Over 35 rare

CNVs were found to encompass 165 genes of possible interest as heterotaxy candidates. Top candidates were screened for L-R patterning functions by morpholino loss of function experiments in Xenopus laevis. We describe results from these analyses and identify the platelet isoform of phosphofructokinase-1 (PFKP) as a novel genetic cause of heterotaxy. Results from these studies collectively confirm a high yield for array-based testing of patients with heterotaxy, and support use of these technologies for identification of novel causative genes.

Previous genetic analyses have suggested that nearly 75% of all X-linked familial and 1% of all sporadic heterotaxy cases can be attributed to mutations in ZIC3. To date, most reported mutations have been identified in five tandemly repeated zinc finger domains in the first two exons. Many of these mutations have demonstrated functional consequences in ZIC3 nuclear transactivation and subcellular localization. The pathogenic potentials of flanking N- and C- terminal mutations are, however, less certain. Therefore, in order to further define the functional significance of mutations occurring throughout the ZIC3 gene, the full ZIC3 coding region and associated splice junctions was sequenced in a cohort of 440 unrelated patients with assorted situs anomalies and CHDs. Of the 11 mutations identified, 8 were novel, including 5 occurring in non-zinc finger domains. For functional studies, we supplemented these 11 mutations with 4 previously reported variants of uncertain significance. Aberrant cytoplasmic shuttling and decreased luciferase reporter transactivation were observed for all mutations affecting zinc finger domains but not for mutations in terminal regions. Results from these analyses significantly expanded the ZIC3 mutation spectrum, supported a higher than expected mutation yield in patients with sporadic heterotaxy (3.8% vs. 1% overall; 5.2% in affected males), and suggested alternative pathogenic mechanisms for mutations affecting non-zinc finger domains.

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DEDICATION:

To my friends and family for their love and support.

To Michael Hester, Kristin Bell, and Thomas Acciani for their Thursdays and Wednesdays.

And to the Prog Gods for all that they do.

I may never find all the answers. I may never understand why. I may never prove what I know to be true, But I know that I still have to try.

Dream Theater Metropolis Pt. 2: Scenes from a Memory

Table of Contents ABSTRACT ...... ii CHAPTER 1: Introduction ...... 1 1.1. THE HETEROTAXY CLINICAL SPECTRUM ...... 1 1.2. HETEROTAXY AND CONGENITAL HEART DISEASE ...... 3 1.3. HETEROTAXY IS GENETICALLY HETEROGENEOUS ...... 4 1.4. GENETIC TESTING FOR HETEROTAXY AND CONGENITAL HEART DEFECTS ...... 7 1.5. ZIC3 AS A GENETIC CAUSE OF HETEROTAXY ...... 10 1.5.1. The ZIC3 mutation spectrum ...... 10 1.5.2. ZIC3 expression and function ...... 12 1.5.3. ZIC3 structure ...... 14 1.6. THE MOLECULAR MECHANISMS OF LEFT-RIGHT PATTERNING ...... 16 1.6.1. The node, cilia, and nodal flow ...... 17 1.6.2. Nodal signaling ...... 22 1.6.3. Flux or flow? ...... 24 1.7. XENOPUS AS A MODEL SYSTEM FOR STUDYING HETEROTAXY ...... 25 1.8. EXPERIMENTAL RATIONALE AND SUMMARY OF MAJOR FINDINGS ...... 26 1.9. FIGURES ...... 29 Figure 1.9.1. ZIC3 structure ...... 29 1.10. REFERENCES ...... 30 CHAPTER 2: Copy number variation as a genetic basis for heterotaxy and heterotaxy-spectrum congenital heart defects ...... 48 2.1. ABSTRACT ...... 48 2.2. INTRODUCTION ...... 49 2.3. MATERIALS AND METHODS ...... 53 2.3.1. Patient recruitment and phenotypic classification ...... 53 2.3.2. Chromosome microarray analysis (CMA) ...... 54 2.3.3. CNV prioritization ...... 55 2.3.4. In vitro fertilization and Xenopus laevis embryo staging ...... 56 2.3.5. Morpholino design ...... 56 2.3.6. pfkp and pitrm1 knockdown ...... 57 2.3.7. Synergy and rescue ...... 58 2.3.8. Left-right marker analyses ...... 59

2.4. RESULTS ...... 60 2.4.1. Genetic analyses of patients with heterotaxy identify rare copy number variants ...... 60 2.4.2. Knockdown of pfkp results in left-right patterning defects in Xenopus laevis ...... 62 2.5. DISCUSSION ...... 65 2.5.1. Copy number variant analyses identify novel heterotaxy gene candidates ...... 65 2.5.2. PFKP as a novel cause of heterotaxy ...... 70 2.5.3. STUDY LIMITATIONS ...... 73 2.6. FUTURE DIRECTIONS ...... 74 2.7. ACKNOWLEDGEMENTS: ...... 75 2.8. TABLES ...... 76 Table 2.8.1. Heterotaxy cohort demographics...... 76 Table 2.8.2. Pathogenic chromosome abnormalities detected in the 225 patient heterotaxy cohort. . 77 Table 2.8.3. Rare CNVs encompassing known heterotaxy genes...... 79 2.9. FIGURES ...... 80 Figure 2.9.1. Overview of CNV findings...... 80 Figure 2.9.2. pfkp but not pitrm1 Xenopus laevis morphants develop organ situs defects by st. 47. .. 81 Figure 2.9.3. Synergy and rescue experiments support MO specificity...... 82 Figure 2.9.4. pfkp knockdown disrupts right-sided coco bias in st. 20-21 embryos...... 83 2.10. SUPPLEMENTARY MATERIALS...... 84 Supp. Figure 2.10.1. Gene ontologies for genes identified in 35 CNVs of interest...... 84 Supp. Table 2.10.1. Phenotypic summary of the 225 patient heterotaxy cohort...... 85 Supp. Table 2.10.2. Rare coding region CNVs detected in the 225 patient heterotaxy cohort...... 87 Supp. Table 2.10.3. Genes identified in rare coding region CNVs detected in the 225 patient heterotaxy cohort...... 92 Supp. Table 2.10.4. Rare copy number variants at loci previously associated with CHD...... 99 Supp. Table 2.10.5. Rare CNVs identified at loci associated with known microdeletion / duplication syndromes but not heterotaxy or CHD...... 101 2.11. REFERENCES ...... 102 CHAPTER 3: Genetic and functional analyses of ZIC3 variants in congenital heart disease ...... 117 3.1. ABSTRACT ...... 117 3.2. INTRODUCTION ...... 118 3.3. MATERIALS AND METHODS ...... 121 3.3.1. Patient recruitment and phenotypic classification ...... 121 3.3.2. Variant detection ...... 122

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3.3.3. Variant interpretation ...... 123 3.3.4. Expression constructs and mutagenesis: ...... 123 3.3.5. Cell culture ...... 123 3.3.6. Luciferase assays...... 123 3.3.7. Immunocytochemistry and subcellular localization ...... 124 3.4. RESULTS ...... 125 3.4.1. Identification of ZIC3 variants ...... 125 3.4.2. ZIC3 variants alter luciferase reporter gene transactivation ...... 128 3.4.3. Zinc-finger domain mutations demonstrate aberrant cytoplasmic subcellular localization .... 129 3.5. DISCUSSION ...... 131 3.6. ACKNOWLEDGEMENTS ...... 137 3.7. TABLES ...... 138 Table 3.7.1. Heterotaxy and CHD cohort demographics...... 138 Table 3.7.2. Summary of ZIC3 variants detected in heterotaxy cohort...... 139 Table 3.7.3. Summary of variant analyses...... 141 3.8. FIGURES ...... 142 Figure 3.8.1. ZIC3 structure...... 142 Figure 3.8.2. Mutations in ZIC3 alter transactivation of an SV40 luciferase reporter...... 143 Figure 3.8.3. Mutations in ZIC3 alter subcellular localization...... 144 3.9. SUPPLEMENTARY MATERIALS...... 145 Supp. Table 3.9.1. Mutagenic primers used to generate HA-tagged variant ZIC3 expression constructs...... 145 Supp. Table 3.9.2. Phenotypic summary of ZIC3 variant carriers relative to the 440 patient heterotaxy and CHD cohort...... 146 3.10. REFERENCES ...... 147 CHAPTER 4: General Discussion ...... 151 4.1. PFKP AS A NOVEL CAUSE OF HETEROTAXY...... 151 4.2. ZIC3 MUTATIONS AS A CAUSE OF HETEROTAXY ...... 155 4.3. ZIC3 AND LEFT-RIGHT PATTERNING ...... 159 4.4. THE FUTURE OF GENETIC TESTING FOR HETEROTAXY ...... 161 4.5. FINAL REMARKS ...... 163 4.6. REFERENCES ...... 165

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CHAPTER 1: Introduction

1.1. THE HETEROTAXY CLINICAL SPECTRUM

Initial establishment of embryonic asymmetry occurs early in development as the embryo defines its anterior-posterior (A-P) and dorsal-ventral (D-V) axes. Left-right (L-R) patterning

(otherwise known as laterality) follows shortly after and manifests most visibly in the asymmetric placement and/or morphogenesis of developing organ systems. In the normal visceral body plan (situs solitus), the heart, stomach, and spleen are located on the left side, while the liver and gallbladder are found on the right. The lungs exhibit side-specific lobation (bi-lobed on the left, tri-lobed on the right). The internal chambers of the heart also possess unique left and right identities and display distinct vessel and valvular connections. Heterotaxy, or situs ambiguus, results when one or more insults in early development alter this stereotypical body plan. It is a multiple congenital anomaly syndrome that lies within a clinical spectrum of laterality disorders ranging from isolated cardiac asymmetries to complete mirroring of internal organ arrangements [situs inversus (SI) totalis]. Disorders of ciliary function, including

Kartagener’s Syndrome (OMIM#244400) and other primary ciliary dyskinesias (PCDs), also lie along this spectrum and reflect a central and conserved role for cilia in L-R patterning (see

Chapter 1.6.1).

Large epidemiological studies of patients with heterotaxy have indicated a population incidence of approximately 1/10,000 (Lin, et al., 2000) with a greater prevalence in Asian (Kim, et al., 2008b) and African American (Correa-Villasenor, et al., 1991) populations. A 2:1 male:female ratio has also been reported (Lin, et al., 2000). Contemporary clinical paradigms consider heterotaxy to be a heterogeneous spectrum of diverse cardiac and visceral abnormalities rather than a defined constellation of pathognomonic defects. In its classic form heterotaxy is

1 characterized by CHDs in combination with anomalies of other visceral organs, such as gut malrotation, abnormalities of spleen position or number, and situs anomalies of the liver and/or stomach. In addition to abnormal positioning and/or formation of these organs, inappropriate retention of symmetric embryonic structures (e.g. persistent left superior vena cava) or loss of normal asymmetry (e.g. right atrial isomerism) can be clues to an underlying laterality disorder.

Abnormalities of the midline, including defects of the ribs and vertebrae, are prevalent and are seen in approximately 40% of all patients (Ticho, et al., 2000). Urinary and genital anomalies, neural tube defects and other midline abnormalities are also common findings (Phoon and Neill,

1994; Bisgrove, et al., 2003). Underscoring this association, the risk for laterality defects has been cited to be three times higher in patients with midline defects than in those without

(Martinez-Frias, 1995; Bisgrove, et al., 2003).

Phenotype-driven classifications have traditionally been used to sub-stratify heterotaxy patients for risk management and a clinical care. When stratified by splenic status, one-year mortality rates have been cited to be as high as 50% (for polysplenia) to 85% (for asplenia)

(Bartz, et al., 2006). Estimates for post-operative mortality in the heterotaxy population as a whole (regardless of splenic status) range from ~20-30% (Serraf, et al., 2010; Sivanandam, et al.,

2014). Most of the associated morbidity and mortality can be attributed to CHDs of varying complexity. Surgical options are diverse and depend on the exact nature of the cardiovascular malformations (reviewed in Kim, 2011). Regardless, they are usually palliative rather than corrective and are typically associated with poorer outcomes and increased mortality relative to patients with equivalent, but isolated, CHDs (Bartz, et al., 2006; Kim, et al., 2008b; Swisher, et al., 2011). Post-operative risks among heterotaxy patients have been found to be especially

pronounced when ciliary dyskinesia is present, owing at least in part to greater incidence of respiratory disease (Jonas, 2011).

1.2. HETEROTAXY AND CONGENITAL HEART DISEASE

Nearly 3% of all CHDs are found in association with laterality disorders (Lin, et al.,

2000; Zhu, et al., 2006). Indeed, most patients with heterotaxy are first brought to clinical attention as a result of neonatal cyanotic CHD (Sutherland and Ware, 2009). We (Ware, et al.,

2004; Cowan, et al., 2014) and others (Megarbane, et al., 2000; De Luca, et al., 2010) have demonstrated that mutations in known heterotaxy genes can also result in isolated CHDs, indicating at least some genetic overlap. It has been estimated that 10% of all patients with heterotaxy will also have a close family history of CHD, with or without accompanying laterality defects (Zhu, et al., 2006). Although no single defect is pathognomonic; particular combinations of CHDs are more frequent, particularly among right or left atrial isomerisms (RAI/LAI)

(reviewed in detail in Lowenthal, et al., 2015). In RAI, right-sided cardiac structures are duplicated at the expense of those on the left, while the opposite is true for LAI. In RAI, common findings include, but are not limited to, absence of the coronary sinus, bilateral superior venae cavae (SVC), complete atrioventricular canal defects (CAVC), ventricular and atrial septal defects (VSD/ASD), pulmonary stenosis/atresia (PS/PA), total or partially anomalous pulmonary venous drainage (TAPVR, PAPVR), atrial and ventricular septal defects (ASDs/VSDs), d- transposition of the great arteries (d-TGA), and single ventricle/hypoplastic left ventricle.

Compared to RAI, LAI-associated CHDs are more variable; however, the inferior vena cava

(IVC) is frequently interrupted. PAPVR, atrioventricular canal defects (AVSD), ASDs/VSDs, and left obstructive defects [coarctation of the aorta (CoA), mitral/aortic stenosis (MS/AS)] are

3 also common findings. Conduction abnormalities can also occur, particularly if the sinus node is duplicated (RAI) or absent (LAI), or if ventricular abnormalities are present.

The enormous heterogeneity of heterotaxy-associated CHDs is arguably best exemplified by recent data released by the National Birth Defects Prevention Study, which describe cardiac and extra-cardiac features of a large cohort of 557 patients with heterotaxy and SI totalis (Lin, et al., 2014). Considering the laterality cohort as a whole, most patients exhibited complex rather than simple CHD (67.7% vs. 9.3%). When the heterotaxy (378 patients) and SI (179 patients) groups were examined independently, most patients with heterotaxy, but not with SI, were found to have complex rather than simple disease (82.8% vs. 7.4% for heterotaxy, 26.6% vs. 14.4% for

SI). Only 9.8% of patients with heterotaxy (vs. 59% of patients with SI) lacked CHDs of any kind. Notably, every major type of CHD was accounted for in the laterality cohort. Several

CHDs [ventricular and atrial septal defects (VSDs, ASDs), double outlet right ventricle (DORV), complete atrioventricular canal defects (CAVC), pulmonary stenosis (PS), pulmonary atresia

(PA) with VSD, total and partial pulmonary venous return (TAPVR, PAPVR), and anomalies of the vena cava] and extra-cardiac anomalies (orofacial clefts, omphalocoeles, and esophageal/bowel atresias) were particularly prevalent among patients with heterotaxy. All told,

~80% of patients with laterality defects exhibited some form of CHD (Lin, et al., 2014).

1.3. HETEROTAXY IS GENETICALLY HETEROGENEOUS

Contemporary advancements in medical care, surgical interventions, and clinical diagnostics have contributed to a well-characterized decrease in patient mortality and concomitant increase in CHD prevalence among patients of reproductive age (Engelfriet, et al.,

2005; Marelli, et al., 2007; Marelli, et al., 2014). Recent analyses indicate that adults now

constitute roughly two-thirds of the CHD population, representing a nearly 60% increase in CHD prevalence among adult patients since the year 2000 (Marelli, et al., 2014). The fact that the greatest increase in CHD prevalence has occurred among the 18- to 40-year-old demographic

(Marelli, et al., 2014) has clear implications for heritability.

With a recurrence odds ratio approaching 80, heterotaxy represents the most highly heritable of all cardiovascular malformations (Oyen, et al., 2009). Although the majority of cases occur sporadically, autosomal recessive, autosomal dominant, and X-linked inheritance patterns have all been reported (Vitale, et al., 2001; Belmont, et al., 2004; Zhu, et al., 2006). In most cases, heterotaxy-associated ciliopathies, such as Kartagener’s syndrome and other forms of

PCD, are inherited in an autosomal recessive manner (for recent reviews see Knowles, et al.,

2013; Kurkowiak, et al., 2015); however, most pathogenic single gene mutations are dominantly inherited (reviewed in Sutherland and Ware, 2009).

Nearly 75% of familial X-linked heterotaxy cases result from loss of function mutations in the zinc finger transcription factor gene, ZIC3 (Ware, et al., 2004; Cowan, et al., 2014).

Despite contributing so strongly to familial X-linked disease, ZIC3 mutations are identifiable in less than 5% of sporadic cases (Ware, et al., 2004; Cowan, et al., 2014). Our lab has used direct sequencing approaches to investigate the genetic contribution of additional genes within the

Nodal signal transduction pathway (NODAL, FOXH1, and CFC1) (unpublished data). Results from these and other studies (Kosaki, et al., 1999a; Kosaki, et al., 1999b; Bamford, et al., 2000;

Karkera, et al., 2007; Selamet Tierney, et al., 2007; Roessler, et al., 2008; Kaasinen, et al., 2010) indicate that mutations affecting Nodal signaling genes, like those impacting ZIC3, account for only a minority (~5-20%) of non-familial heterotaxy cases (reviewed in Sutherland and Ware,

2009). Signaling pathway mutations are most frequently detected in NODAL (~5-10%) and

CFC1 (6-21%), while LEFTYA, GDF1, and ACVR2B are more rarely affected (1-3%). Pathway relevant mutations in the Smad-interacting transcription factor, FOXH1, have also been reported

(Roessler, et al., 2008). Mutation yields for these genes are surprisingly low considering the central role played by the Nodal pathway in L-R patterning. Because animal models homozygous for Nodal signaling mutations fail to properly form mesoderm and arrest during gastrulation

(Peeters and Devriendt, 2006), it has been proposed that the relative scarcity of Nodal signaling mutations in heterotaxy patients may partially reflect high lethality among affected embryos.

Accordingly, laterality defects would only be expected in patients with abrogated Nodal signaling that remained at levels still permissive for progression through the earliest stages of embryonic development.

In addition to mutations in known heterotaxy genes, L-R patterning defects have also been reported in association with a large number of genetic syndromes, including the aforementioned ciliopathies. Diverse chromosomal causes of heterotaxy have additionally been identified and range in severity from whole chromosome aneuploidies (trisomy 13, trisomy 18) to smaller chromosomal rearrangements (inversions, unbalanced translocations) and copy number variation (CNVs) in the form of macro, micro, and submicroscopic deletions and duplications (Bisgrove, et al., 2003; Song, et al., 2009; Fakhro, et al., 2011; Rigler, et al., 2014).

As a group, these CNVs can be broadly defined as structural genetic variation that results in either an increase or a decrease in the number of copies of a particular segment of DNA.

Investigations completed to date indicate that CNVs may explain roughly 20-30% of heterotaxy cases (Fakhro, et al., 2011; Rigler, et al., 2014). It also appears likely that at least a subset of heterotaxy cases can arise from combinatorial interaction of multiple susceptibility or modifying loci. Involvement of a second causative locus has been suggested as an explanation for situs

abnormalities in patients with heterozygous mutations in ACVR2B (Kosaki, et al., 1999b) and

LEFTYA (Kosaki, et al., 1999a). Compound heterozygosity of NODAL and HNF3 mutations has also been reported in a patient with situs abnormalities (Kosaki, et al., 1999b), echoing phenotypic observations from Nodal/Hnf3 double heterozygous mice (Collignon, et al., 1996).

Interactions between susceptibility loci may ultimately help to explain tendencies toward high phenotypic diversity and variable penetrance among related members of affected families

(Chung, et al., 2011). Collectively, these genetically diverse causes underscore the enormous complexity of human heterotaxy and suggest that a large proportion of causative factors remain to be determined. Supporting this assertion, ongoing animal studies (predominantly completed in mice, chick, zebrafish, and frog) have already identified more than 80 genes with involvement in

L-R patterning pathways, many of which have yet to be associated with human disease (Rigler, et al., 2014).

1.4. GENETIC TESTING FOR HETEROTAXY AND CONGENITAL HEART DEFECTS

Over the last two decades, development of cutting edge genetic testing technologies has advanced at an unprecedented pace. Novel sequencing and chromosome-based methods have improved the speed and breadth of available clinical testing options and have provided scientists with exciting new tools for genetic research and disease profiling. Although chromosome analysis (karyotyping) remains the gold standard for clinical diagnosis of many aneuploidies and other large chromosomal abnormalities, fluorescence in situ hybridization (FISH) and chromosomal microarray (CMA) technologies have proven to be invaluable for identifying microdeletion and duplication syndromes resulting from abnormalities too small to be detected by conventional chromosomal analyses. In practice, the relatively limited resolutions of

7 karyotyping and FISH have rendered them insufficient to identify a genetic cause in the majority of patients with CHDs of unknown etiology (Breckpot, et al., 2010) and in nearly half of all patients with syndromic CHD (Breckpot, et al., 2011). Consequently, CMA has emerged as a high fidelity alternative for first-line CHD genetic testing, particularly when extra-cardiac features are present but a suspected diagnosis is lacking (Breckpot, et al., 2011).

Early heterotaxy genetic studies, such as those leading to the identification of ZIC3

(Casey, et al., 1993; Ferrero, et al., 1997), relied heavily on cytogenetic linkage analyses to define broad regions of genetic interest. These pioneering studies were subject to many technical and practical limitations, including low resolution, high testing costs, and strict requirements for large pedigrees with multiple affected family members. Universal adoption of polymerase chain reaction (PCR) and introduction of automated sequencing technologies helped to facilitate gene mapping on a finer scale while more broadly paving the way for the Human Genome Project and other large-scale efforts to characterize the human genome. Since that time, heterotaxy gene discovery has been greatly assisted by application of these technologies to the study of early L-R patterning in diverse model systems (Bisgrove, et al., 2003; Blum, et al., 2009). Experiments in these organisms have helped to identify the transforming growth factor-beta (TGF- ligand,

Nodal, as the core signaling molecule governing laterality across vertebrate species and have provided a strong impetus for sequencing studies targeting critical Nodal pathway elements

(Kosaki, et al., 1999a; Kosaki, et al., 1999b; Bamford, et al., 2000; Goldmuntz, et al., 2002;

Selamet Tierney, et al., 2007; Roessler, et al., 2008; Mohapatra, et al., 2009). (see Chapter 1.6.2 for further discussion of Nodal signaling). As a direct result of these and other sequencing efforts

(Britz-Cunningham, et al., 1995; Bartoloni, et al., 2002; Olbrich, et al., 2002; Robinson, et al.,

2003; Ware, et al., 2004; Hornef, et al., 2006; Karkera, et al., 2007; Schwabe, et al., 2008; Failly,

et al., 2009; Maitra, et al., 2010; Tariq, et al., 2011; Li, et al., 2012; Perles, et al., 2012; Saunders, et al., 2012; Cowan, et al., 2014), clinical heterotaxy panels now routinely offer targeted gene sequencing for nearly 20 genes, many of which are components of the Nodal pathway (reviewed in Sutherland and Ware, 2009).

As many of these gene sequencing efforts were underway, comparative genomic hybridization (CGH), which was first introduced in the early 1990s as a way to detect chromosomal abnormalities in tumor cells (Kallioniemi, et al., 1992), was undergoing a similar technological renaissance. Innovative application of array technology to CGH protocols (array-

CGH) helped to expand CNV analyses to a previously unprecedented scale and opened the door for genome-wide assessment of patients with diverse genetic diseases (reviewed in Vissers, et al.,

2005; Bejjani and Shaffer, 2006; Carter, 2007). Limitations in the ability of array-CGH to detect copy-neutral cytogenetic abnormalities (uniparental disomy, loss of heterozygosity, balanced translocations), as well as low copy repeats and segmental duplications have been at least partially overcome by combined use of array-CGH and higher resolution single nucleotide polymorphism (SNP) genotyping platforms (Haraksingh, et al., 2011; Wiszniewska, et al., 2014).

These advancements have helped to improve resolution and maximize potential for CNV detection, particularly in the study of developmental delay and mental retardation (Friedman, et al., 2009; Bernardini, et al., 2010; Fan, et al., 2013). As previously mentioned, application of these CMA technologies to the study of heterotaxy and/or CHD has been enormously fruitful, identifying CNVs as a significant genetic cause in up to 30% of patients and leading to the identification of several novel heterotaxy genes (Fakhro, et al., 2011; Lander and Ware, 2014;

Rigler, et al., 2014).

Although not a focus of this thesis, it is important to recognize the emergence of massively parallel next-generation sequencing (NGS) technologies and their growing application to the study of genetic disease. These commercially and methodologically distinct services share similar foundations in repeated sequencing of DNA fragments and assembly of the resulting reads into a completed sequence (reviewed in Dorn, et al., 2014, with a focus on CHD). Higher depths of coverage, faster turnaround times, and increased cost-effectiveness are possible when compared to traditional capillary-based sequencing methods (Shendure and Ji, 2008; Mardis,

2011). To date, few CHD studies and even fewer analyses of heterotaxy patients have been completed using NGS technologies (Tariq, et al., 2011; Arrington, et al., 2012; Al Turki, et al.,

2014; Francis, et al., 2014; Zaidi, et al., 2013; reviewed in Rabbani, et al., 2014). In 2011, we reported the first (and to date only) novel heterotaxy gene to be identified by whole exome sequencing (WES): the cell shape regulator, SHROOM3 (Tariq, et al., 2011). Homozygous missense mutations were identified in a single patient with heterotaxy and in 4/96 patients tested in follow-up. These results supported a role for SHROOM3 in L-R patterning, potentially through interaction with ROCK2, a known regulator of laterality and recognized SHROOM3 binding partner (Fakhro, et al., 2011; Wang, et al., 2011). It is anticipated that continued application of high resolution microarray and emerging NGS testing methods will lead to identification of many new and promising heterotaxy candidate genes.

1.5. ZIC3 AS A GENETIC CAUSE OF HETEROTAXY

1.5.1. The ZIC3 mutation spectrum

Casey et al. (Casey, et al., 1993) provided the first genetic evidence supporting existence of a heterotaxy locus on the X-chromosome. These initial analyses incorporated linkage data

from 39 members of a large family with severe X-linked disease and broadly localized the critical disease interval to Xq24-q27.1. Subsequent fine mapping of members of the same family, as well as identification of a similarly localized submicroscopic deletion in an unrelated male patient, helped to narrow the critical region to a 1.3Mb stretch at Xq26.2 (Ferrero, et al., 1997).

Landmark experiments building on these initial studies precisely mapped ZIC3 to this locus and demonstrated for the first time that mutations in ZIC3 can be causative for X-linked heterotaxy

(Gebbia, et al., 1997).

Since that time, over 20 novel ZIC3 mutations have been identified in patients with heterotaxy and/or CHDs (Gebbia, et al., 1997; Megarbane, et al., 2000; Ware, et al., 2004; Fritz, et al., 2005; Tzschach, et al., 2006; Chhin, et al., 2007; De Luca, et al., 2010; Wessels, et al.,

2010; Chung, et al., 2011; D'Alessandro, et al., 2011; Ma, et al., 2012; Cowan, et al., 2014). As is true for most X-linked diseases, it is not uncommon for males with hemizygous ZIC3 mutations to exhibit more severe phenotypes than heterozygous female carriers (Bisgrove, et al., 2003).

Nevertheless, affected females have been frequently reported (Ferrero, et al., 1997; Ware, et al.,

2004; Fritz, et al., 2005; Tzschach, et al., 2006; Chhin, et al., 2007; Ma, et al., 2012). Indeed, my own recent analysis of ZIC3 mutations in a large cohort of 440 patients identified variants in

5/181 (~3%) tested female patients (Cowan, et al., 2014, included here as Chapter 3). At least four of these variants were considered to be causative for disease based on described functional analyses. The overall proportion of penetrant disease in female mutation carriers that can be attributed to skewed X-inactivation remains uncertain as results from inactivation studies have only rarely been reported (Chhin, et al., 2007; Ma, et al., 2012).

Recent analyses of ZIC3 mutations in patients with heterotaxy have highlighted an intriguing phenotypic overlap with VACTERL syndrome (vertebral anomalies, anal atresia,

11 cardiovascular malformations, tracheoesophageal fistula, renal anomalies, limb abnormalities), suggesting potential for shared developmental etiologies (Wessels, et al., 2010; Chung, et al.,

2011; D'Alessandro, et al., 2013; Cowan, et al., 2014). Notably, one of these patients was found to carry a two-alanine expansion of the ZIC3 polyalanine tract (Wessels, et al., 2010). Although its functional significance was untested, the absence of the mutation in parental or 192 ethnically matched controls led the authors to conclude that it likely contributed to the patient’s abnormal phenotype. Similar polyalanine tract expansions were subsequently identified in other patients with heterotaxy and/or CHD (D'Alessandro, et al., 2013, Cowan, et al., 2014). Collectively, these studies revealed a wider clinical spectrum for heterotaxy mutations than previously appreciated and supported inclusion of patients with extra-cardiac malformations who might otherwise lack overt laterality defects in future heterotaxy genetic studies.

1.5.2. ZIC3 expression and function

A total of five ZIC (Zinc Finger in the Cerebellum) genes have been identified in humans and mice (Aruga, et al., 1996a; Aruga, et al., 1996b; Furushima, et al., 2000). The ZIC genes derive their name from shared expression in the adult mouse cerebellum and from common function as developmentally significant zinc finger transcription factors. During mouse embryogenesis, expression of each Zic gene demonstrates a unique and dynamic expression pattern (Nagai, et al., 1997; Elms, et al., 2004; Inoue, et al., 2004). Pre-gastrulation, Zic3 is restricted to ectodermal epiblast, but as gastrulation begins, expression shifts to include the primitive streak, the mesodermal “wings” of the embryo, and the adjacent ectoderm (Nagai, et al., 1997; Elms, et al., 2004). By onset of neurulation there is an increasing trend towards dorsal ectodermal expression. Zic3 is notably absent from the node during the earliest stages of

primitive streak formation, but becomes strongly expressed there at least as early as E7.75 (Elms, et al., 2004; Sutherland, et al., 2013). As development continues, Zic3 can be readily detected in limb bud mesenchyme and in the rhombomeres of the developing neural tube (Nagai, et al.,

1997). Expression in the heart has been more controversial as it has been detected solely by

PCR-based methods (RT-PCR, qPCR) and, even then, only at low levels (Purandare, et al., 2002;

Zhu, et al., 2007b; Bedard, et al., 2011; Haaning, et al., 2013). Nevertheless, the most pertinent question is not what quantity of Zic3 is expressed in the embryonic heart, but whether cardiac expression of ZIC3 is important for normal heart development. In agreement with other published studies (Purandare, et al., 2002; Bedard, et al., 2007), our lab was unable to detect cardiac ZIC3 expression by WISH in recent analyses of multiple Zic3-LacZ-BAC transgenic lines (Haaning, et al., 2013; Sutherland, et al., 2013). We considered this lack of expression alongside other available evidence - normal cardiac marker expression (Tbx5, Nppa), normal myocardial trabeculation, and newly recognized roles for ZIC3 in node morphogenesis (Jiang, et al., 2013; Sutherland, et al., 2013) - and concluded that the extensive heart defects observed in

ZIC3 null and hypomorphic mice are most likely a consequence of earlier defects in L-R patterning rather than a primary insult on the developing heart.

These broad expression patterns hint at the diversity of ZIC3 embryonic function. In addition to well-documented roles in L-R patterning (Kitaguchi, et al., 2000; Purandare, et al.,

2002; Ware, et al., 2006a; Ware, et al., 2006b; Zhu, et al., 2007b; Zhu, et al., 2007a; Jiang, et al.,

2013; Sutherland, et al., 2013), ZIC3 is important for neural and neural crest development

(Nakata, et al., 1997; Klootwijk, et al., 2000), limb bud digitation (Quinn, et al., 2012), gastrulation (Ware, et al., 2006b; Cast, et al., 2012), and stem cell pluripotency (Lim, et al.,

2007). Recent data from Xenopus also indicates involvement in Wnt pathways, specifically in

13 suppression of-catenin signaling during notochord and organizer formation (Fujimi, et al.,

2012). Within the context of L-R patterning, ZIC3 has been shown to act upstream of Nodal signaling (Purandare, et al., 2002; Ware, et al., 2006a) and to be required for morphogenesis of the embryonic node (Jiang, et al., 2013; Sutherland, et al., 2013). More in-depth discussion of node formation and its role in Nodal signaling can be found in Chapter 1.6. For further discussion of ZIC3 function in node morphogenesis, refer to Chapter 4.3.

1.5.3. ZIC3 structure

Until recently, all five ZIC genes were thought to consist of three exons, with the first exon-intron boundary being fully conserved among all family members and the second exon- intron boundary being present in only ZIC1, ZIC2 and ZIC3 (Grinberg and Millen, 2005). In

2011, our lab identified a novel alternatively spliced ZIC3 transcript (ZIC3-B) that preferentially splices in a fourth exon at the expense of the normally terminal third exon (Bedard, et al., 2011).

The first 407 amino acid residues of the ZIC3-A and ZIC3-B protein products are therefore identical, with the only sequence differences manifesting in the C-terminal ends of each isoform.

To date, only two sequencing efforts have included exon 4; however, no mutations were identified amongst the 109 (Bedard, et al., 2011) or 440 (Cowan, et al., 2014) heterotaxy patients tested. Expression analyses revealed that both isoforms exhibit similar subcellular and embryonic expression patterns with only marginal differences in relative abundance, although only ZIC3-A was detectable in the embryonic mouse heart (Bedard, et al., 2011). Initial studies suggest the potential for isoform-specific functions, as only ZIC3-A was capable of synergistically activating a glioblastoma (Gli) binding site reporter when co-transfected with Gli3. Additional experiments

will be required to determine the precise role(s) played by ZIC3-B and the functional significance, if any, of the fourth exon.

The ZIC genes share significant homology with the Drosophila odd-paired (OPA) and cubitus interruptus (Ci) genes, as well as with the mammalian GLI family. Proteins encoded by these genes all contain a series of C2H2 zinc finger domains, each of which comprises pairs of cysteine and histidine residues arranged in tetrahedral array around a lone zinc ion. Positioning of individual zinc fingers in tandem confers highly stable attachment to transcriptional targets

(Iuchi, 2001). In ZIC3, five of these fingers span the first two exons (Figure 1.9.1) and are essential for proper transcriptional activation (Ware, et al., 2004; Cowan, et al., 2014).

Importantly, nuclear localization (NLS) and nuclear export (NES) signals have been found to overlap these regions (Bedard, et al., 2007; Hatayama, et al., 2008) and reflect a growing recognition of ZIC3 as a protein capable of nuclear-cytoplasmic shuttling. The NLS is interspersed in structure, occupying amino acid residues that span all five zinc finger domains

(292-356, 367-382, 403-412) (Bedard, et al., 2007; Hatayama, et al., 2008). In contrast, the NES is an isolated signal located at the boundary of the second and third zinc fingers (positions 313-

325) (Bedard, et al., 2007). This export signal is usually hidden and requires either a normal or pathological change in protein structure to be exposed (Bedard, et al., 2011). Unsurprisingly, mutations affecting the NES (Bedard, et al., 2007), as well as those affecting the interspersed

NLS (Ware, et al., 2004; Chhin, et al., 2007; Hatayama, et al., 2008; Cowan, et al., 2014), consistently disrupt subcellular localization of ZIC3 protein, resulting in abnormal numbers of cells with predominantly nuclear or cytoplasmic ZIC3. Transcriptional activities are also typically compromised by these mutations, as demonstrated by altered trans-activation of in vitro luciferase reporters. The finding that the zinc finger domains, the NLS, and the NES occupy

15 overlapping sequences is not unprecedented, as this arrangement has been noted among other transcription factors, including members of the related GLI superfamily (LaCasse and Lefebvre,

1995; Black, et al., 2001; Fernandez-Martinez, et al., 2003; Xiao, et al., 2003).

Relative to mutations affecting zinc finger regions, mutations impacting upstream and downstream N- and C-terminal regions have been identified with less frequency. We recently reported genetic and functional analysis of a large subset of 11 ZIC3 variants (including 5 in the

N-terminal region, as well as the most C-terminal variant identified to date) in Cowan et al.

(Cowan, et al., 2014), which is included here as Chapter 3. The reader is referred there for experimental details and in-depth analyses; however, a major takeaway is that terminal mutations do not appear to impact ZIC3 transcriptional activation or subcellular localization (Cowan, et al.,

2014, Chapter 3). Additional studies will be required to discern the pathogenic potential of the

N- and C-terminal regions and the functional impact of mutations in these regions.

1.6. THE MOLECULAR MECHANISMS OF LEFT-RIGHT PATTERNING

The preceding sections clearly illustrate that heterotaxy is a disease of considerable clinical and genetic heterogeneity and not one that can be simply explained by mutations in a single gene or pathway. Established and emerging genome-wide testing methods hold great potential for identifying novel heterotaxy genes with myriad developmental functions. As candidate genes are newly identified, it will be important to consider their functions within the holistic context of known laterality programs.

Current understanding of the molecular mechanisms governing L-R patterning recognizes a number of distinct developmental paradigms. The prevailing “nodal flow” model proposes that laterality is established shortly after gastrulation by asymmetric, left-sided activation of Nodal

signaling. These signaling asymmetries are instigated by extracellular fluid flow over a transient, ciliated organizer located at the posterior end of the notochord. This organizer is alternatively termed the embryonic node in mice, the posterior notochordal plate (PNC) in rabbit, Hensen’s node in chick, Kupffer’s vesicle (KV) in zebrafish, or the gastrocoel roof plate (GRP) in frog

(Sulik, et al., 1994; Nonaka, et al., 1998; Essner, et al., 2005; Schweickert, et al., 2007; Vick, et al., 2009). Although cilia are notably absent in the chick node (Gros, et al., 2009), the importance of nodal flow in Xenopus and zebrafish has been demonstrated (Essner, et al., 2002; Essner, et al., 2005; Schweickert, et al., 2007). Such a high degree of interspecies conservation of function supports a strongly conserved role for nodal flow in many vertebrate models. Nevertheless, accumulating evidence, particularly in Xenopus, indicates that some species may establish even earlier asymmetries by restricting ion transporters to right-sided blastomeres (Levin, et al., 2002;

Levin, 2004; Fukumoto, et al., 2005b; Fukumoto, et al., 2005a; Adams, et al., 2006; Aw, et al.,

2008; Morokuma, et al., 2008; Aw, et al., 2010; Vandenberg, et al., 2013). These cleavage-stage ionic asymmetries are thought to drive similar asymmetric redistribution of serotonin, itself a functionally critical L-R patterning signal (Fukumoto, et al., 2005b; Fukumoto, et al., 2005a).

The extent to which early (ion-flux) and late (nodal flow) L-R patterning pathways interact remains extremely controversial and, on the whole, poorly understood (Vandenberg and Levin,

2010; Schweickert, et al., 2012).

1.6.1. The node, cilia, and nodal flow

Evidence for the involvement of cilia in L-R patterning first emerged in 1976 with report of dynein arm abnormalities and immotile cilia in patients with Kartagener’s syndrome

(Afzelius, 1976). Although Rott had speculated as early as 1980 that abnormal cilia motility may

17 underlie pathogenesis of Kartagener’s syndrome (Rott, 1980), it would be nearly 20 years before cilia were identified on the surface of the node and their developmental link to laterality was proposed (Sulik, et al., 1994). In 1997, mutations in the ciliary molecular motor, left-right-dynein

(Lrd), were identified as being causative for laterality defects occurring in the well-characterized iv (inversus viscerum) mutant mouse line (Supp, et al., 1997). Shortly thereafter, landmark experiments in knockout mice lacking the microtubular motor protein, Kif3B, provided the first definitive evidence that L-R patterning is driven by motile cilia on cells of the embryonic node and that these cilia generate an asymmetric, left-directed extracellular fluid flow across the entire surface of the node (Nonaka, et al., 1998). These and other pioneering studies solidified node monocilia as important structures for L-R patterning and singled out dynein and kinesin molecular motors as critical drivers of node cilia motility (Okada, et al., 1999; Takeda, et al.,

1999; Essner, et al., 2002; McGrath, et al., 2003; Essner, et al., 2005; Kramer-Zucker, et al.,

2005).

Structurally and functionally homologous structures (collectively: left-right organizers,

LROs) have since been identified in a number of diverse species including frog (Schweickert, et al., 2007), zebrafish (Essner, et al., 2005), medaka (Okada, et al., 2005a), and rabbit (Okada, et al., 2005a; Blum, et al., 2007), indicating a high degree of interspecies conservation of cilia- based L-R patterning mechanisms. In mouse, the node is the last of three transient organizers to develop during primitive streak formation (Kinder, et al., 2001). Each of these organizers exhibits similar inductive potential but differs in relative gene expression profile and cell lineage contributions (Beddington, 1994; Sulik, et al., 1994; Tam, et al., 1997; Kinder, et al., 2001). The node’s role as a developmental organizer was definitively demonstrated by an elegant series of transplantation experiments that illustrated the potential for ectopic node tissue to induce novel

neural axes in late-stage gastrulas and to subsequently promote tissue differentiation appropriate for the ectopic axial plan (Beddington, 1994; Tam, et al., 1997). Follow-up ablation experiments confirmed a necessary function for the node in patterning the L-R axis (Davidson, et al., 1999).

The critical stages of mouse node morphogenesis occur shortly after gastrulation (~E7-

E8). At this time, endodermal cells on the presumptive node surface progressively give way to an underlying sheet of ciliated cells at the extreme edge of the posterior notochordal plate

(Yamanaka, et al., 2007; Lee and Anderson, 2008). The mechanism regulating removal of the overlying endodermal cells remains uncertain; however, the number of exposed ciliated cells is known to progressively increase throughout node morphogenesis (Lee and Anderson, 2008), culminating in formation of a concave, tear-drop shaped structure comprised of ~200-300 ciliated cells. The equivalent LRO structure in the frog, the GRP, also forms at the start of neurulation (stage 13) and is derived from a superficial layer of mesoderm that involutes through the dorsal blastopore lip during gastrulation (Shook, et al., 2004; Schweickert, et al., 2007). The

GRP subsequently takes shape as a triangle of superficial mesoderm flanked by larger endodermal cell populations (lateral endodermal crests, LECs) (Shook, et al., 2004). Ciliation first becomes apparent by stage 14 and reaches a peak at stage 16 with ~250-270 monociliated cells. Both cilia length and nodal flow velocity increase throughout these stages, leveling off at

~5m and 4m/s respectively by stage 18-19. Developing motile cilia in both mouse and frog progressively orient and tilt posteriorly to better generate a unidirectional (rather than vortical) fluid flow (Nonaka, et al., 2005; Okada, et al., 2005b; Schweickert, et al., 2007). This positioning is governed by planar cell polarity (PCP) pathways, which, in general, act as important regulators of global tissue orientation by coordinating localized cellular polarities (Hong and Dawid, 2009;

Maisonneuve, et al., 2009; Antic, et al., 2010; Borovina, et al., 2010; Song, et al., 2010; Gray, et

19 al., 2011; Wallingford and Mitchell, 2011; Gegg, et al., 2014). In the case of cilia orientation at the LRO, the membrane protein VANGL2 has been identified as the core PCP protein involved

(Antic, et al., 2010; Borovina, et al., 2010; Song, et al., 2010).

Two ciliated cell populations have been identified at the mouse node, including a central population of “pit cells” that contain motile, flow-generating cilia and a peripheral population of

“crown cells” containing predominantly non-motile, sensory cilia (McGrath, et al., 2003;

Yoshiba, et al., 2012). Initial structural analyses suggested that the rotational motility of node cilia may result from absence of a central pair of singlet microtubules that are typically observed in other motile cilia populations (Bellomo, et al., 1996). This 9+0 arrangement (which references the 9 peripherally arranged doublet microtubule pairs) is more commonly observed in non-motile sensory cilia, including those adorning crown cells in the node periphery. Nevertheless, recent identification of motile cilia with a traditional 9+2 arrangement in both the zebrafish KV

(Kramer-Zucker, et al., 2005) and mouse node (Caspary, et al., 2007) indicate that not all motile cilia at the LRO lack a central pair. Furthermore, mouse or zebrafish models harboring mutations in the central pair genes, Hydin, Spag6, and rsph9 do not display L-R patterning defects (Sapiro, et al., 2002; Lechtreck, et al., 2008; Castleman, et al., 2009; Olbrich, et al., 2012) and situs abnormalities are not typically observed among patients with PCD caused by central pair defects

(Tamalet, et al., 2001; Stannard, et al., 2004; Castleman, et al., 2009; Zietkiewicz, et al., 2012), suggesting a lack of involvement of the central pair in L-R patterning. For the moment, the functional distinction between 9+0 and 9+2 motile cilia populations remains uncertain, as do the exact mechanisms dictating ciliary rotational movement.

At least in mouse, frog, and zebrafish, a key genetic determinant of node ciliogenesis is the forkhead transcription factor, Foxj1, which is thought to be the master regulator (Chen, et al.,

1998; Blatt, et al., 1999; Brody, et al., 2000; Stubbs, et al., 2008; Yu, et al., 2008). In mouse, the homeobox transcription factor, Noto, is also critically important for both node formation and associated ciliogenesis and appears to operate independently of fibroblast growth factor (FGF) signaling (Beckers, et al., 2007; Alten, et al., 2012). Cilia development and maintenance is driven by a complex intraflagellar transport (IFT) system, which functions to transport molecular cargo up and down the length of axoneme (reviewed in Bhogaraju, et al., 2013). Core IFT proteins, including the aforementioned kinesin and dynein molecular motors, have been identified to be important for not only L-R patterning (Supp, et al., 1997; Nonaka, et al., 1998;

Marszalek, et al., 1999; Takeda, et al., 1999; Rana, et al., 2004; Hong and Dawid, 2009), but also for transduction of developmentally critical signals, including Sonic Hedgehog (Shh) (Huangfu, et al., 2003; Huangfu and Anderson, 2005; Liu, et al., 2005; Huangfu and Anderson, 2006) and

Wingless (Wnt) (Corbit, et al., 2008; Jopling and Izpisua Belmonte, 2009). These are far from being the only L-R relevant signals: Bone morphogenic protein (Bmp) (Schilling, et al., 1999;

Piedra and Ros, 2002; Furtado, et al., 2008; Veerkamp, et al., 2013), Notch (Krebs, et al., 2003;

Lopes, et al., 2010; Kato, 2011), and Retinoic Acid (RA) pathways (Chazaud, et al., 1999;

Kawakami, et al., 2005) have all been variously identified as important contributors to laterality.

Most significant, however, is the Nodal signaling cascade, which forms the central backbone to all current L-R patterning paradigms and represents a major focus of past and present genetic testing efforts (Kosaki, et al., 1999a; Kosaki, et al., 1999b; Bamford, et al., 2000; Goldmuntz, et al., 2002; Selamet Tierney, et al., 2007; Roessler, et al., 2008; Mohapatra, et al., 2009).

1.6.2. Nodal signaling

One of the first major breakthroughs in L-R patterning was the discovery of left-sided

Nodal signaling at the lateral plate and left-sided Shh signaling at Hensen’s node in early chick embryos (Levin, et al., 1995). Similar left-sided expression of Nodal was soon identified in the mouse, alongside asymmetric expression of additional Nodal pathway components (eg: Lefty,

Pitx2) (Collignon, et al., 1996; Lowe, et al., 1996; Meno, et al., 1996). Extracellular flow generated at the node was subsequently reported as the cause of asymmetric Nodal activation in the left lateral plate (Okada, et al., 1999), providing the missing link between observed signaling asymmetries and demonstrations of cilia-driven flow at the node.

Nodal is initially expressed symmetrically and only later becomes restricted to the left side during flow stages. Interestingly, Nodal induces not only its own expression but also that of its monomeric repressor, Lefty2 (reviewed in Hamada, et al., 2002). The resulting signaling gradients establish feedback loops that encourage left-sided expression and right-sided repression of Nodal (Sakuma, et al., 2002; Muller, et al., 2012). The end result is asymmetric activation of important downstream Nodal signaling components, including the homeodomain protein, Pitx2, a key regulator of left-sided organogenesis (Logan, et al., 1998; Yoshioka, et al., 1998; Shiratori, et al., 2001).

A second molecular asymmetry both precedes and feeds into these Nodal signaling pathways. This conserved gene (a BMP, TGF-, and Wnt pathway inhibitor variously known as coco in frogs, charon in fish, and Dante/Cerl2 in mammals; henceforth referred to as coco) is initially bilaterally co-expressed alongside Nodal in the lateral plate (Hashimoto, et al., 2004;

Schweickert, et al., 2010). During post-flow stages, however, coco mRNA becomes increasingly concentrated on the right side. Because Coco functions as a potent Nodal inhibitor, these

expression biases are essential for establishing the left-sided Nodal signaling cascade required for downstream asymmetric organogenesis (Marques, et al., 2004; Hojo, et al., 2007; Vonica and

Brivanlou, 2007; Schweickert, et al., 2010; Tingler, et al., 2014). Supporting this critical role, genetic manipulation of coco expression by morpholino knockdown (on the right side) or nucleotide microinjections (on the left) have been shown to result in L-R patterning defects in

70-100% of treated embryos (Vonica and Brivanlou, 2007; Schweickert, et al., 2010; Tingler, et al., 2014; reviewed in Blum, et al., 2014). Elucidation of this core Nodal signaling pathway provided for the first time a logical and experimentally supported mechanism for laterality determination. However, many important questions still remained unanswered: by what mechanism does nodal flow initiate asymmetric left-sided Nodal signaling? What are the relevant molecular signals? And are events at the node the initiating step in L-R patterning or does an earlier mechanism set the stage for LRO-stage asymmetries?

Two models emerged to attempt to provide an explanation as to how nodal flow might initiate the left-sided Nodal signaling cascade. The first model hypothesized that the directional fluid flow generated by ciliary motion transports one or more critical morphogens from the node to the left peri-nodal region by an undetermined transport mechanism. Shh and RA were suggested as plausible morphogen candidates since both are known to be transported from the node in nodal vesicular parcels (NVPs) in response to FGF signaling (Tanaka, et al., 2005). An alternative “two-cilia” model was proposed as an attempt to reconcile theoretical and experimental discrepancies associated with the “morphogen transport” theory (reviewed in Tabin and Vogan, 2003). According to this model, immotile sensory cilia of the node periphery act as antennae that recognize leftward flow generated by motile cilia of central pit cells. The mechano- sensory stimulation resulting from nodal flow was hypothesized to activate a calcium-mediated

23 signal response required for activation of Nodal signaling in the left-lateral plate mesoderm

(Tabin and Vogan, 2003). Support for the two-cilia model has been bolstered by evidence from mutant mice deficient in Pkd2, a calcium ion channel that has been previously linked to autosomal dominant polycystic kidney disease (ADPKD) and to sensation of flow-induced mechanical stress in renal cilia (with Pkd1) (Nauli, et al., 2003). Pkd2 is expressed in both motile and immotile node cilia and Pkd2-dependent calcium signaling is known to be asymmetrically elevated in the left peri-nodal region (McGrath, et al., 2003). Importantly, mouse studies have demonstrated a requirement for peri-nodal Pkd2-mediated calcium signaling for proper L-R patterning (Yoshiba, et al., 2012). Pkd2 is thought to act through interaction with Pkd1l1, a node- specific paralogue of renal Pkd1 (Field, et al., 2011). Given the already established mechano- sensory function for Pkd2 in renal cells, these results appear to validate the two-cilia model.

Nevertheless, because a transported morphogen could theoretically activate calcium signaling in a Pkd2-dependent fashion, the morphogen transport model cannot be fully discounted.

1.6.3. Flux or flow?

Although nodal flow and asymmetric Nodal signaling appear to be highly conserved mechanisms for L-R patterning, it is not yet clear whether events at the LRO represent the initiating steps of the laterality program. An alternative cleavage-stage mechanism has been proposed based on data generated from experiments in frog, zebrafish, and chick. This “early” model hypothesizes that asymmetric localization of ion-transporters during initial cleavages are thought to generate ionic and membrane voltage gradients across individual blastomeres. These gradients are then proposed to drive asymmetric localization of charged molecules important for

L-R patterning by way of gap junction communication between neighboring cells (Levin, et al.,

2002; Levin, 2004; Fukumoto, et al., 2005b; Fukumoto, et al., 2005a; Adams, et al., 2006; Aw, et al., 2008; Morokuma, et al., 2008; Aw, et al., 2010; Vandenberg, et al., 2013). Much focus has been placed on serotonin as a critical “early” determinant as it has been reported to not only asymmetrically localize to the right ventral blastomere in response to established ionic and voltage gradients, but also to be functionally critical for initiating proper organ situs and L-R relevant signaling in frog and chick embryos (Fukumoto, et al., 2005b; Fukumoto, et al., 2005a;

Vandenberg, et al., 2013).

It is important to recognize that the “early” (ion flux) and “late” (nodal flow) models are not necessarily mutually exclusive, particularly as cleavage-stage perturbations in ion-flux and serotonin signaling have been shown to be capable of dysregulating Nodal signaling pathways

(Fukumoto, et al., 2005a). Nevertheless, the question as to what, if any, molecular steps may bridge earlier cleavage and later node-stage asymmetries remains a point of considerable debate and one of the major open questions in contemporary L-R patterning studies (Beyer, et al., 2012;

Vandenberg, et al., 2013; Blum, et al., 2014; Tingler, et al., 2014). For further discussion of this important topic, the reader is referred to Chapter 4.1.

1.7. XENOPUS AS A MODEL SYSTEM FOR STUDYING HETEROTAXY

Animal models have long served as vital tools for understanding key events in axial patterning, dating back at least as far as Spemann’s groundbreaking early experiments in amphibians (Spemann and Falkenberg, 1919; Spemann and Mangold, 1924). In recent years, the

African clawed frog, Xenopus, has emerged as a particularly versatile system for the study of laterality (reviewed in Blum, et al., 2009). Among other advantages, the relatively large size of

Xenopus eggs facilitates a wide array of embryonic manipulations, while the large quantity of

25 oocytes obtained from hormone-injected females provides an easily renewable pool of embryos for experimentation. Understanding of vertebrate L-R patterning has benefited greatly from experiments in Xenopus, particularly following identification of the GRP as a bona-fide LRO

(Schweickert, et al., 2007). Ionic and molecular asymmetries preceding GRP formation have also been identified and have opened additional avenues for experimentation (Levin, et al., 2002;

Levin, 2004; Fukumoto, et al., 2005b; Fukumoto, et al., 2005a; Adams, et al., 2006; Aw, et al.,

2008; Morokuma, et al., 2008; Aw, et al., 2010; Vandenberg, et al., 2013) (see Chapter 1.5.2). In both frog and zebrafish, targeted morpholinos knockdowns have been used extensively for identifying novel heterotaxy candidates and for investigating gene function in L-R pattering pathways (Stubbs, et al., 2008; Fakhro, et al., 2011; Cast, et al., 2012). These referenced studies are not comprehensive, but do provide a sampling of the many reports published in the two decades since morpholino technologies were first introduced (Partridge, et al., 1996; Summerton and Weller, 1997). Emerging CRISPR/Cas9 genome editing systems are already beginning to revolutionize knockdown studies in zebrafish, and hold similar potential for gene screening and functional testing in Xenopus (Blitz, et al., 2013; Nakayama, et al., 2013)

1.8. EXPERIMENTAL RATIONALE AND SUMMARY OF MAJOR FINDINGS

The overall goal of this dissertation has been to further delineate the genetic causes and developmental mechanisms underlying heterotaxy and CHD. Mutations in ZIC3 are the most common genetic cause of heterotaxy in families with X-linked disease and have been previously estimated to explain approximately 1% of all sporadic heterotaxy cases (Ware, et al., 2004).

Nevertheless, point mutations in genes currently linked to pathways governing early L-R

patterning are detected in only a small proportion of heterotaxy patients. With such genetic heterogeneity, there remains enormous potential for novel gene discovery.

Work in this thesis therefore aimed to balance identification and developmental characterization of novel heterotaxy genes with investigations into the genetic mechanisms of

ZIC3-related heterotaxy. Our lab sought to identify novel L-R patterning genes by screening a large, 225 patient heterotaxy cohort for genome-wide copy number variation. Using array-based

CGH and SNP genotyping methods, we identified rare CNVs with pathogenic potential in 46 patients, representing a total yield of 20.4% for the cohort. These CNVs encompassed genes with diverse biological functions, including genes already recognized to cause laterality defects in human patients and animal studies, as well as novel heterotaxy candidates. Follow-up functional validation in Xenopus laevis suggested a critical role for PFKP, the platelet isoform of phosphofructokinase-1, in L-R patterning. Overall, our results supported the use of genome-wide

CNV analyses in detecting novel genetic causes of heterotaxy and identified a subset of promising candidate genes for future developmental study.

To investigate the functional significance of novel ZIC3 mutations detected in patients with heterotaxy, our lab sequenced complete ZIC3 coding regions and splice junctions of 440 unrelated patients with assorted situs abnormalities and CHDs. Using cell culture systems, we assessed the impact of detected mutations on reliable readouts of ZIC3 function. Our results greatly expanded the known ZIC3 mutation spectrum (particularly in poorly characterized N- and

C-terminal regions), revealed a higher than anticipated mutation prevalence in sporadic heterotaxy, supported past genotype-phenotype correlations between mutations in zinc finger regions and laterality defects, confirmed the importance of these zinc finger domains for ZIC3

27 trafficking and function, and indicated that mutations in N- and C-terminal domains may follow pathogenic mechanisms distinct from those affecting zinc finger domains.

1.9. FIGURES

Figure 1.9.1. ZIC3 structure. Wild-type human ZIC3, RefSeq: NP_003404.1, NM_003413.3. cDNA numbering begins at the A (position +1) of the ATG initiation codon (codon 1). NLS = nuclear localization signal, NES = nuclear export signal, PolyAla = polyalanine tract, UTR = untranslated region, ZF = zinc-finger domain

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CHAPTER 2: Copy number variation as a genetic basis for heterotaxy and heterotaxy- spectrum congenital heart defects

2.1. ABSTRACT

Genomic disorders and rare copy number abnormalities are identified in 15-25% of patients with syndromic conditions, but their prevalence in individuals with isolated birth defects is less clear. Occurring in 8 per 1000 live births, congenital heart defects (CHDs) are the most common birth defect and a significant source of worldwide morbidity and mortality. A spectrum of CHDs is seen in heterotaxy, a highly heritable and genetically heterogeneous multiple congenital anomaly syndrome resulting from failure to properly establish left-right (L-R) organ asymmetry during early embryonic development. To identify novel genetic causes of heterotaxy, we analyzed copy number variants (CNVs) in 225 patients with heterotaxy and heterotaxy- spectrum CHDs using complementary array-CGH and SNP-array genotyping methods. Identified abnormalities ranged from large unbalanced translocations to smaller, kilobase-scale CNVs, including a rare, single exon deletion in ZIC3. CNVs were identified in ~20% of patients and encompassed both known and putative heterotaxy genes. Morpholino loss-of-function experiments in Xenopus support a role for one of these novel candidates, the platelet isoform of phosphofructokinase-1 (PFKP) in heterotaxy. Collectively, our results confirm a high CNV yield for array-based testing in patients with heterotaxy, and support use of CNV analysis for identification of novel biological processes relevant to human laterality.

2.2. INTRODUCTION

Heterotaxy is a relatively infrequent (~1:10,000) multiple congenital anomaly syndrome resulting from abnormal specification of the left-right (L-R) body axis during early embryonic development (Lin, et al., 2000). It is considered to exist along a clinical spectrum of laterality disorders ranging from isolated dextrocardia to situs inversus (SI) totalis that also includes left and right isomerisms. In its classic form heterotaxy is characterized by combined occurrence of visceral situs abnormalities (gut malrotations, stomach and liver anomalies, abnormalities of spleen positioning or number) and congenital heart defects (CHDs) of varying complexity, which account for the majority of associated morbidity and mortality. Over 96% of patients with heterotaxy exhibit some form of CHD (Lin, et al., 2014), often requiring surgical intervention.

Clinical outcomes are disproportionately poorer than in patients without heterotaxy who have similar cardiac defects and are typified by prolonged courses and significantly greater likelihood for post-surgical complications (Bartz, et al., 2006; Kim, et al., 2008b; Swisher, et al., 2011).

This clinical picture firmly establishes heterotaxy as not only a disease of significant phenotypic heterogeneity, but also one of considerable medical and economic consequence.

Although heterotaxy typically occurs sporadically and with unknown cause, heritability is highest among all classes of CHDs, supporting existence of a strong genetic component. (Oyen, et al., 2009). Autosomal recessive, autosomal dominant, and X-linked inheritance patterns have all been described (Vitale, et al., 2001; Belmont, et al., 2004; Zhu, et al., 2006). Mutations in the zinc finger of the cerebellum 3 (ZIC3) gene are particularly well-documented (Gebbia, et al.,

1997; Ware, et al., 2004; D'Alessandro, et al., 2011; D'Alessandro, et al., 2013; Cowan, et al.,

2014) and are considered to be causative in the majority (~75%) of familial X-linked pedigrees.

Surprisingly, however, ZIC3 mutations underlie only a minority (3-5%) of sporadic heterotaxy

49 cases (Ware, et al., 2004; Cowan, et al., 2014). Likewise, despite a conserved and central role for

Nodal signaling in establishment of early molecular asymmetries (Sulik, et al., 1994; Nonaka, et al., 1998; Essner, et al., 2005; Schweickert, et al., 2007; Vick, et al., 2009), point mutations in

Nodal pathway components are also not routinely identified and collectively explain only 5-10% of heterotaxy cases (Roessler, et al., 2008; Mohapatra, et al., 2009). Mutations in other causative genes are detected at similar or even lower frequencies (reviewed in Sutherland, et al., 2013), indicating significant genetic heterogeneity. As a specific genetic etiology is currently identifiable in only a minority of patients (Sutherland, et al., 2013), there remains enormous potential for novel gene and pathway discovery.

Copy number variants (CNVs) in the form of complex chromosomal rearrangements and submicroscopic duplications and deletions have increasingly become recognized as important causes of birth defects and neurodevelopmental disease (Pinto, et al., 2010; Coe, et al., 2014;

Hooli, et al., 2014; Rees, et al., 2014; Vondrackova, et al., 2014). Depending on size and genomic position, CNVs can encompass complete or partial intronic or exonic regions, can include one or multiple genes, or can disrupt regulator regions such as promoters and enhancers.

If a CNV shifts the normal reading frame, this change may lead to premature truncation or loss- of-function through nonsense-mediated decay (NMD) (Nguyen, et al., 2014; Valdivielso-Ramos, et al., 2014) or gain-of-function if the transcript escapes decay (Ben-Shachar, et al., 2009). Gene interruptions and fusions, positional effects, recessive mutation unmasking, and allelic transvection events are all additional potential molecular consequences (reviewed in Zhang, et al., 2009). The pathogenic significance of a particular CNV is, therefore, highly dependent on the location of its breakpoints, the genic content of the intervening deleted or duplicated segment, and the genomic landscape in which the CNV is situated.

Studies of patients with CHD indicate that CNVs are a major genetic cause of cardiovascular disease, occurring in 3-25% of patients with extra-cardiac abnormalities and in 3-

10% with isolated heart defects (reviewed in Lander and Ware, 2014). To date, two studies have employed genome-wide approaches to explicitly examine the role of CNVs in heterotaxy

(Fakhro, et al., 2011; Rigler, et al., 2014). Analyzing a cohort of 262 patients with classic heterotaxy and/or isolated heterotaxy spectrum CHDs, Fakhro et al. (Fakhro, et al., 2011) identified 45 rare CNVs in 39 patients, representing a 15% CNV yield in their phenotypically mixed population. A slightly higher rate of 26% (20 CNVs in 19/74 patients) was reported in a more recent study that restricted analysis solely to patients with classic heterotaxy (Rigler, et al.,

2014). Yields from both of these studies approximated those seen in other CHD cohorts

(reviewed in Lander and Ware, 2014) and together support CNVs as important contributor to human laterality.

It is important to recognize that not all CNVs are pathogenic and that copy number variation is detected even amongst healthy individuals. A recent meta-analysis of 55 CNV studies culled from the Database of Genomic Variants (DGV) (dgv.tcag.ca/) indicates that CNVs are likely to occupy between 4.8-9.5% of the human genome (Zarrei, et al., 2015).

Approximately 5-10% of the general population is estimated to carry at least one CNV spanning less than 500kb, with an even greater proportion of individuals (~65-80%) having a CNV less than 100kb (Mikhail, 2014). In order to systematically catalogue this genetic variation, online and clinical databases, such as the DGV, have been established as repositories of commonly occurring CNVs. These databases serve as vital reference points for clinical and research efforts focused on identifying CNVs of pathogenic significance to genetic disease.

The flexibility and versatility of array comparative genomic hybridization (aCGH), and single-nucleotide polymorphism (SNP) microarrays have made them excellent tools for both gene discovery and clinical practice (Thienpont, et al., 2007; Richards, et al., 2008; Breckpot, et al., 2010; Goldmuntz, et al., 2011; Baker, et al., 2012; Connor, et al., 2014). In the present study, we have screened a large cohort of patients with heterotaxy spectrum malformations using two complementary methods: (1) a custom-designed whole genome aCGH array (Boone, et al., 2010) and (2) a non-targeted, genome-wide SNP array with comparatively even marker spacing. We hypothesized that use of both platforms would maximize ability to detect disease-relevant CNVs and to identify genes with previously unrecognized roles in L-R patterning. In total, we identified rare CNVs in 46/225 (20.44%) patients ranging in size from large, megabase-scale translocations to smaller, kilobase-scale CNVs. Excluding CNVs that were considered to be clearly pathogenic due to their overall size and complexity, or their genomic content (encompassing regions or genes previously associated with or identified as definitive causes of heterotaxy/CHD), we reduced the number of CNVs suitable for candidate gene analysis to 35 in 30/225 (13.33%) patients. Morpholino-based loss-of-function screens in Xenopus laevis were then used to confirm roles for gene candidates in L-R patterning. In this report, we describe overall findings from these CNV screens while highlighting functional testing results for the platelet isoform of phosphofructokinase 1 (pfkp), which we have identified as a novel genetic cause of heterotaxy.

Collectively, our analyses support CNVs as a significant contributor to heterotaxy causation and reiterate the value of genome-wide CNV screening as a tool for identifying novel laterality genes and pathways.

2.3. MATERIALS AND METHODS

2.3.1. Patient recruitment and phenotypic classification

All studies were approved by the Institutional Review Board at Cincinnati Children’s

Hospital Medical Center (CCHMC). Detailed phenotypic information was collected from patient histories and from chart review, with emphasis placed on thoracic and abdominal situs, splenic status, presence or absence of gall bladder abnormalities, imperforate anus, and cardiac structure.

Patients were classified as having SI totalis, heterotaxy, or isolated heterotaxy spectrum CHD following previously defined criteria (Ware, et al., 2004). As some heterotaxy patients with ZIC3 mutations also exhibit VACTERL-like (vertebral anomalies, anal atresia, cardiovascular malformations, tracheoesophageal fistula, renal anomalies, limb abnormalities) phenotypes, patients with these features are also noted. Disease was considered to be “familial” if (1) the presenting pedigree demonstrated autosomal dominant, autosomal recessive, or X-linked inheritance, (2) there was more than one family member with heterotaxy or laterality disorder regardless of their degree of relationship to the proband, (3) there was heterotaxy in the proband and a first degree relative with isolated CHD, or (4) there was heterotaxy in the proband and a first degree relative with situs-related defects. All other patients were considered to have sporadic disease. Patients with previously identified cytogenetic findings or with mutations in known heterotaxy genes were excluded from further analysis. No patient had a previously identified CNV from prior clinical screening. In total, the final cohort comprised 225 unrelated patients with assorted situs and/or cardiac abnormalities, including 139 males and 86 females.

Full cohort demographics are summarized in Table 2.8.1.

2.3.2. Chromosome microarray analysis (CMA)

Blood samples were collected, and genomic DNA was prepared following standard protocols. Prior to testing, personal health information was removed from each patient and a unique identification number was assigned. To evaluate for CNVs, samples were genotyped using two complementary CMA methods: genome-wide SNP analysis and array comparative genomic hybridization (aCGH). We hypothesized that use of both array types would provide a greater breadth of genomic coverage for detection of disease-relevant CNVs, while also validating CNVs identified on both platforms. The specific platform(s) used for each patient was dictated by sample availability at time of testing.

SNP genotyping was performed using the Illumina Human370-DNA Analysis BeadChip

(42 patients) or Illumina HumanOmni1-Quad Beadchip (106 patients) platforms (Illumina Inc.,

San Diego, CA, USA). These chips encompassed approximately 370,000 SNP markers (mean spacing 7.7kb, median 5kb) or 1,000,000 SNP markers (mean spacing 2.4kb, median 1.2kb) respectively. The Illumina Infinium Assay was performed as described by the manufacturer using 250 ng of patient DNA. B-allele frequency and log2 R ratio were analyzed with Illumina

GenomeStudio V2009.2 software and DNA copy number changes were prioritized using output from cnvPartition Plug-in v2.3.4 software.

Array-CGH experiments were completed using a custom array designed by Baylor

Medical Genetics Laboratories (221 patients) [http://www.bcm.edu/geneticlabs/cma/tables.html]

(Ou, et al., 2008). This array has been previously described (Boone, et al., 2010) and was designed to encompass approximately 180,000 oligonucleotides with whole genome coverage of

30 kb resolution, and exon by exon coverage of over 1,700 genes (mean spacing

4.2probes/exon). Generated data were normalized using the Agilent Feature Extraction Software.

Copy number segmentation was accomplished using two algorithms: (1) a segmentation method using circular binary segmentation with additional post-processing steps to remove spurious calls and filter for high evidence calls; and (2) the Biodiscovery’s rank segmentation algorithm, which uses a parametric model rather than permutation testing. QC scores were calculated for data qualities of all arrays.

Genotyping data from both platforms were independently analyzed and interpreted by two board-certified cytogeneticists. Spurious calls were eliminated from raw data and CNVs meeting criteria for clinical relevancy were determined following accepted standards for clinical reporting at each institution. Throughout this report, we provide results derived from each platform separately whenever possible. For CNVs identified by aCGH, size data are presented as minimum and maximum values, with gene content determined using maximum predicted breakpoints. Breakpoint predictions for all CNVs were completed using human reference sequence Build 36.1, hg18.

2.3.3. CNV prioritization

CNVs were prioritized for candidate gene analysis based on the following criteria: (1) absence of the CNV in online (Database of Genomic Variants, http://dgv.tcag.ca/) and clinical

(CCHMC, Baylor) databases of human genetic variation; (2) published expression of affected genes in L-R critical developmental stages (data from both Xenopus and mouse studies were considered, with expression in Xenopus laevis emphasized); (3) number of affected genes in the

CNV interval (priority was given to CNVs encompassing between 1 and 6 genes to best facilitate systematic candidate gene screening), and (4) number of potentially pathogenic CNVs in the affected patient (patients with fewer CNVs were prioritized).

2.3.4. In vitro fertilization and Xenopus laevis embryo staging

Sexually mature wild-type male and female Xenopus laevis were purchased from Nasco.

Oocyte collection and in vitro fertilization protocols were performed as previously described

(Cast, et al., 2012). For all experiments, collected embryos were staged according to the Xenopus developmental table (Nieuwkoop and Faber, 1994).

2.3.5. Morpholino design

All morpholinos were obtained through GeneTools, LLC. Translation-blocking (TB) morpholinos targeting the translation start-site were designed for pfkp (5’-

CTTCCCGAGCCCTCTCTCTACTCTC-3’) and pitrm1 (5’-

TCTGCCTCACAGCCGCACAACTATC-3’) using NCBI reference sequences

NM_001097850.1 and NM_001086512, respectively. A splice-blocking (SB) morpholino targeting the pfkp exon 8-intron 8 splice junction (5’-AATGAACACAAGCCACTGTACCTCA-

3’) was also generated using splice junction sequence obtained by PCR amplification of stage 47

Xenopus laevis genomic DNA (sense 5’-TGTTTATCCCTGAATACCC-3’; antisense 5’-

CAAACTCCCCTAAACAAGTTA-3’). A fluorescein-conjugated zic3 morpholino (5’-

ACAATAAAACTTACCTTCATGTGCT-3’) targeting the exon 2-intron 2 splice junction was used as a positive control. A standard negative control morpholino (5’-

CCTCTTACCTCAGTTACAATTTATA-3’) was purchased to control for effects of microinjection.

2.3.6. pfkp and pitrm1 knockdown

Expression of pfkp during L-R critical stages was confirmed by RT-PCR using the following cDNA primers: sense: 5’-AACCTGTTCCGGGAAGAATGGAGT-3’, anti-sense 5’-

ACCCACAGTGTCTTCCCATGACTT-3’. ODC (ornithine decarboxylase) primers (sense 5′-

GCCATTGTGAAGACTCTCTCCATTC-3′, antisense 5′-TTCGGGTGATTCCTTGCCAC-3′) were used as a PCR amplification control (Heasman, et al., 2000; Cast, et al., 2012). The expression profile of pitrm1 was not examined as organ situs defects were not significantly increased over uninjected and control MO injected control embryos (see Chapter 2.4.2). All morpholinos were injected into 2-cell stage Xenopus laevis embryos suspended in 4% Ficoll in

1/3 X Marc’s modified Ringer’s (MMR) buffer. Injections were targeted to the animal poles of both blastomeres. For organ situs assessments, embryos were cultured in 0.1X MMR until stage

47, at which time they were anesthetized in 0.05% benzocaine and scored for heart, gut, and gallbladder positioning. In wild-type embryos at this stage, the heart should be centrally located with a leftward-looping outflow tract, the gut should originate on the right and coil in a counter- clockwise direction, and the gallbladder should be right-sided. Deviations from these expectations were considered to be abnormal. Embryos with abnormal situs of two or more organs were classified as having heterotaxy, while embryos with involvement of only a single organ system were considered to have an isolated situs anomaly. Embryos with mirror-image reversal of all three organs were given a classification of SI. Morpholinos were titrated to the lowest dosages required to yield organ situs defects [pfkp TB = 1.25ng / 3nL injection, pfkp SB =

25ng / 3nL injection, zic3 SB - 5.3ng / 3nL injection]. Pitrm1 morpholinos were tested across a range of 1ng-5ng / 3nL injection. Efficiency of pfkp splice blockade was determined at two stages by RT-PCR. Total RNA was extracted from stage 9 (pre-gastrulation) and stage 25

(tailbud stage) Xenopus laevis embryos by standard TRIzol methods (Life Technologies). Exon 8 deletion was confirmed by gel electrophoresis and bi-directional sequencing of the deletion product using primers targeting the 3’ end of exon 7

(5’CCAGAGGACCTTTGTGCTGGAAGTTATG-3’) and the 5’ end of exon 9 (5’-

ACACAACACAGGCTGGCGTTTC-3’). ODC primers were again used for control of RNA quality and PCR amplification.

2.3.7. Synergy and rescue

Morpholino synergy and mRNA rescue experiments were performed to confirm specificity of targeting. For synergy experiments, sub-threshold dosages of pfkp TB (0.83ng) and

SB (16.7ng) morpholinos were co- or independently injected into both animal poles of 2-cell stage embryos. Morpholino synergy was demonstrated by a significant increase in organ situs defects in co-injected embryos relative to independently injected embryos at stage 47. For mRNA rescue experiments, in vitro transcribed full-length human PFKP mRNA was co-injected with pfkp SB morpholino into both animal poles of 2-cell stage embryos. Successful rescue was demonstrated by a significant decrease in organ situs defects in mRNA-treated embryos relative to untreated morphants. Full-length human PFKP cDNA for mRNA synthesis was generated from Origene cDNA clone SC118507 by PCR amplification (sense 5’-

GCAGAGCTCGTTTAGTGAACCGC-3’, antisense 5-GATGGGCACTCGCCGATTAG-3’).

Identities of the cDNA restriction fragment and PCR product were both confirmed by Sanger sequencing. PFKP mRNA was subsequently in vitro transcribed by T7 polymerase using the mMessage mMachine kit (Ambion) following manufacturer’s protocols and stored at -80°C. For rescue experiments, embryos were injected with either 500pg or 1000pg of transcribed mRNA,

either alone or in combination with pfkp MO titrations sufficient to cause laterality defects in morphant embryos.

2.3.8. Left-right marker analyses

Xenopus laevis embryos were fixed overnight at 4°C in MEMFA (4% paraformaldehyde in MEM salts). Fixed embryos were subsequently dehydrated through three consecutive 5 minute

100% methanol washes and stored at -20°C. An antisense RNA probe for the L-R marker, coco was generated from a pCMV-SPORT6 plasmid (provided by Dr. Mustafa Khokha) using a

T7/SP6 DIG RNA Labeling Kit (Roche) following manufacturer’s instructions. Whole-mount in situ hybridizations (WISH) were performed as previously described (Sive, et al., 2000), with one major modification: prior to hybridization, fixed stage 20-21 embryos were bisected along the transverse plane into separate anterior and posterior halves, briefly post-fixed (10 minutes in 4%

MEMFA), and run through a graded methanol series for storage. To prevent accidental loss of signal-containing fragments, hybridization was performed on both halves, which were then sorted and appropriately trimmed. Staining was accomplished using BM Purple alkaline phosphatase chromogenic substrate (Roche). Images were captured using a Nikon SMZ1500 stereomicroscope outfitted with a Nikon DXM1200F digital camera and processed using Nikon

Act-1 (v. 2.62) imaging software. Coco expression was scored as left-sided biased, right-sided biased, or right/left unbiased by visual inspection. Final embryo counts included only those embryos that could be unequivocally classified into one of the three distinct categories.

2.4. RESULTS

2.4.1. Genetic analyses of patients with heterotaxy identify rare copy number variants

In this study, we utilized complementary genotyping methods – an exon-targeted custom aCGH oligo array (Agilent, Boone, et al., 2010) and a set of untargeted genomic SNP arrays

(Illumina) - to investigate copy number variation in a large cohort of 225 patients with heterotaxy and heterotaxy spectrum malformations. The major cardiovascular and laterality phenotypes of all patients are summarized in Supp. Table 2.10.1. Of the 225 patients tested, 148

(65.78%) were genotyped by SNP array, 221 (98.22%) were genotyped by aCGH, and 144

(64.00%) were genotyped using both platforms. To maximize our chances of identifying novel causes of heterotaxy, we focused our attention on rare CNVs encompassing coding regions of one or more genes. In total, we identified novel pathogenic or likely pathogenic CNVs in 46/225 patients, representing an overall CNV yield of 20.4% (Figure 2.9.1). Thirteen patients had large/complex chromosomal abnormalities (summarized in Table 2.8.2), which notably included a ~3Mb deletion affecting the 22q11.2 critical region identified in a female patient with abdominal situs abnormalities and CHD (Patient 6). Another three CNVs affected genes previously associated with heterotaxy (Table 2.8.3): a 553-686kb 2p25.1 duplication encompassing ROCK2 in a male patient with sporadic and complex CHD (Patient 15), a 1.47Mb-

1.61Mb 3p24.1 deletion encompassing TGFBR2 in a female patient with familial heterotaxy

(Patient 16) and a 2.9-3.3kb Xq26.2 deletion encompassing the last exon of ZIC3 in a male patient with sporadic heterotaxy (Patient 14). The latter CNV was particularly noteworthy as

ZIC3 deletions have only rarely been reported (Chung, et al., 2011).

To best facilitate identification of novel heterotaxy genes, smaller CNVs with lower gene content and without known causation for heterotaxy or CHD (henceforth “CNVs of interest”)

were prioritized. Twenty-two (62.86%) of the 35 CNVs meeting these criteria involved gains of genetic material, while 13 (37.14%) involved genetic losses (Supp. Table 2.10.2). As an estimation of CNV gene content, we performed breakpoint predictions using available marker/probe positioning. These analyses were inherently limited by marker and probe spacing, but suggested that 1/22 (4.55%) of the detected gains and none of the detected losses were likely to have at least one breakpoint in exonic sequence. Similarly, 15/22 (68%) gains and 5/13

(38.46%) losses were predicted to encompass the entire coding region of at least one gene. Only one of the 22 identified gains (4.55%) and four (58%) of the 13 identified losses (30.77%) were predicted to be lie entirely within a single gene. These CNVs ranged in size from 114-350kb

(mean: 250.36kb; median: 286.74kb) on SNP array and 10-427kb (mean: 202.74-259.89kb; median: 234.94-301.31kb) by aCGH. The remaining CNVs affected partial coding sequences or coding sequences of more than one gene. There was no obvious chromosome bias among detected CNVs, with CNVs of interest detected on 21/23 chromosomes – the exceptions being the X chromosome and chromosome 21 (data not shown).

Of the 35 CNVs of interest, 7 were >1Mb in size (SNP size range 1.01-2.78Mb, mean

1.96Mb, median 2.06Mb; aCGH size range 1.08-3.84Mb, mean 2.02-2.30Mb, median 1.86-

2.58Mb), and encompassed between 1-18 genes (mean 7; median 7). The remaining 27 CNVs individually affected <1Mb of sequence (SNP size range 161.66-877.08kb, mean 458.50kb, median 472.45kb; aCGH size range 4.90- 960.81 kb, mean 322.04-387.01kb, median 205.73-

279.31kb) impacting between 1 and 16 genes each (mean 3.96; median 2).

A total of 165 genes with diverse developmental and molecular functions were encompassed by the 35 CNVs of interest (Supp. Table 2.10.3, Supp. Figure 2.10.1). Among these genes were a subset with recognized roles in developmental processes and pathways

61 important for L-R patterning, including ciliogenesis, transforming growth factor- (TGF) signaling, and cell-cell communication. Eleven of the 35 CNVs of interest encompassed loci previously associated with CHD (Supp. Table 2.10.4), while one (a 1.70-2.59Mb 2q13 deletion identified in a patient with sporadic heterotaxy) disrupted a block of ten genes previously deleted in a similarly affected heterotaxy patient (Rudd, et al., 2009). CNVs encompassing loci associated with known microduplication / deletion syndromes but not with heterotaxy or CHD are summarized in Supp. Table 2.10.5. For candidate gene screens we prioritized analysis of genes with known expression during critical L-R developmental stages in animal studies. Priority was given to genes encompassed by CNVs with lower gene content and identified in patients with fewer CNVs of interest so as to better facilitate functional studies.

2.4.2. Knockdown of pfkp results in left-right patterning defects in Xenopus laevis

Because L-R patterning processes are highly conserved across vertebrate species (Sulik, et al., 1994; Nonaka, et al., 1998; Essner, et al., 2005; Schweickert, et al., 2007; Vick, et al.,

2009), significant knowledge regarding human laterality can be gleaned from animal studies The merits of Xenopus as model for laterality are extensive and have been reviewed in detail elsewhere (Blum, et al., 2009). Importantly, a large number of embryos can be obtained through in vitro fertilization with negligible effort, providing a renewable resource for in vivo genetic and developmental manipulations. We therefore undertook a Xenopus-based morpholino loss of function approach to screen identified CNVs for potential heterotaxy candidate genes. This method not only provided a more efficient method for gene screening than costly and time- consuming mouse-modeling studies, but was also already of proven utility, having been

previously used to successfully identify a number of novel heterotaxy genes (Fakhro, et al.,

2011).

A single patient from the heterotaxy cohort was found to carry a heterozygous 175kb deletion, encompassing the PFKP and PITRM1 genes, which encode the platelet isoform of phosphofructokinase-1 (PFK-1) and the protease pitrilysin metallopeptidase 1, respectively

(Figure 2.9.2, panel a). This CNV was of interest as it was identified using both genotyping platforms, was not described in online databases of genomic variation, had not been seen in institutional cytogenetic lab database, and contained an easily testable number of genes, one of which (PFKP) is a recognized interaction partner of the H+-V-ATPase proton pump (Su, et al.,

2003; Su, et al., 2008), a known regulator of left-right patterning (Adams, et al., 2006). To assess the effect of pfkp and pitrm1 knockdown on L-R patterning, TB and SB morpholinos

(GeneTools, LLC.) were designed and injected into both blastomeres of 2-cell stage Xenopus laevis embryos. Gut, heart, and gallbladder were then scored for abnormal situs at stage 47 according to published criteria (Branford, et al., 2000). A significantly greater proportion of pfkp, but not pitrm1, morphants developed organ situs defects relative to uninjected or control morpholino-injected embryos (Figure 2.9.2, panel c). Organ situs defects in pfkp morphants were dose-dependent, with greater proportions of embryos exhibiting disrupted laterality at higher morpholino concentrations (data not shown). Consistent with previous reports demonstrating

PFK-1 expression in early development (Davidson, et al., 1983; Raddatz and Lovtrup-Rein,

1986; Dworkin and Dworkin-Rastl, 1991), pfkp was expressed throughout stages known to be important for L-R patterning in Xenopus (Figure 2.9.2, panel b). These results indicated that pfkp but not pitrm1 was a suitable candidate for further study.

In order to confirm pfkp mRNA knockdown in pfkp splice block morphants RT-PCR and gene-specific amplification of whole RNA from st. 9 (not shown) and st. 25 embryos (Figure

2.9.2, panel d, lane 2) was completed. Sequencing of the deletion product revealed activation of a cryptic splice site 13bp downstream of the intron 7-exon 8 junction, but confirmed deletion of the majority of exon 8 (96/109bp). The resulting pfkp transcript is predicted to encode a truncated, non-frameshifted 754aa (vs. 786aa) protein lacking residues predicted to contribute to

ATP binding. Synergy and rescue experiments using wild-type human PFKP mRNA supported specificity of the splice-block morpholino effect (Figure 2.9.3).

In order to confirm an effect for pfkp knockdown on L-R relevant signaling pathways, we next examined expression of the conserved L-R patterning marker, coco, at the gastrocoel roof plate (GRP). Ciliated cells of the GRP function analogously to those of the mouse embryonic node, generating a highly directional extracellular fluid flow that asymmetrically activates Nodal signaling pathways in the left lateral plate (Schweickert, et al., 2007). In this setting, coco serves as a potent inhibitor of the Nodal ligand, becoming increasingly restricted to the right side of the

GRP and functioning to prevent right-sided Nodal expression during flow relevant stages

(Rodriguez Esteban, et al., 1999; Yokouchi, et al., 1999; Hashimoto, et al., 2004; Marques, et al.,

2004; Hojo, et al., 2007; Vonica and Brivanlou, 2007; Schweickert, et al., 2010). This right-sided bias is of critical importance for the laterality program: without it, Nodal signaling asymmetries are improperly established and resulting embryos develop extensive laterality defects (Vonica and Brivanlou, 2007; Schweickert, et al., 2010; Tingler, et al., 2014; reviewed in Blum, et al.,

2014). Consistent with previous reports (Schweickert, et al., 2010), we observed a right-sided coco expression bias in ~60% of untreated late-flow stage embryos (stage 20-21) (Figure 2.9.4).

This bias was significantly reduced in pfkp morphants (44.8%, p = 0.0058). Supporting

functional impact, the magnitude of this reduction was comparable to that observed following knockdown of dnah9, which encodes an axonemal molecular motor essential for GRP ciliary motility and extracellular fluid flow (Schweickert, et al., 2010). On the basis of these morpholino studies, we identify pfkp as a novel heterotaxy gene and conclude that the 175kb CNV encompassing this gene is the most likely explanation for the laterality phenotype observed in our patient.

2.5. DISCUSSION

2.5.1. Copy number variant analyses identify novel heterotaxy gene candidates

As the majority of heterotaxy cases remain unexplained, the primary goal of our study was to identify novel genetic variation of pathogenic significance to this patient population. To this end, we have performed comprehensive genome-wide CNV analyses on a large cohort of

225 patients with heterotaxy and CHD. In contrast to previous analyses (Fakhro, et al., 2011), patients were carefully phenotyped, permitting distinction between patients with classic heterotaxy situs abnormalities and patients with isolated CHD. Excluding CNVs of prohibitively large gene content (Table 2.8.2) and those encompassing genes previously identified as causes of heterotaxy (Table 2.8.3), we detected 35 unique CNVs of potential interest for functional testing.

Collectively, these variants encompassed 165 genes of diverse biological and molecular ontology

(Supp. Table 2.8.3, Supp. Figure 2.10.1), including a subset with roles in pathways and processes relevant to laterality. These genes are individually described below.

The “nodal flow” model of L-R patterning proposes that the vertebrate laterality program is initiated by asymmetric, activation of Nodal signaling in the left lateral plate shortly after gastrulation. Evidence from multiple species has solidified the importance of transient, ciliated

65 organizers in establishing and propagating these signaling asymmetries (Sulik, et al., 1994;

Nonaka, et al., 1998; Essner, et al., 2005; Schweickert, et al., 2007; Vick, et al., 2009). Mutations affecting ciliary genes have subsequently been identified as a major cause of heterotaxy spectrum disorders ranging from “classic” heterotaxy (CHD with visceral situs abnormalities) to primary ciliary dyskinesia (PCD) and other heterotaxy-spectrum ciliopathies (reviewed in

Sutherland and Ware, 2009). There is significant overlap between PCD and heterotaxy with 6-

12% of PCD patients having heterotaxy (Kennedy, et al., 2007; Shapiro, et al., 2014) and an unknown but significant percentage of heterotaxy patients having PCD (Shapiro, et al., 2014).

Results from our CNV screens have identified rare variants encompassing a number of genes with cilia-related functions, including TTC21B, CEP290, TTBK2, and CFAP126. Two of these,

CEP290 and TTC21B, are recognized causes of human ciliopathies and are known to play critical roles in cilia assembly and transport (Tran, et al., 2008; Coppieters, et al., 2010).

Mutations in TTC21B, which encodes the retrograde intraflagellar transport protein,

THM1/IFT139, cause both nephronophthisis 12 (end-stage renal disease) and short-rib thoracic dysplasia 4 (skeletal anomalies with or without polydactyly) (Davis, et al., 2011). Similarly,

TTC21B knockdown in mice results in the “alien” phenotype, which is characterized by preaxial polydactyly, cleft palate, micro-ophthalmia, and brain abnormalities. Retrograde ciliary transport is disrupted, leading to impaired cilia growth and downstream Sonic hedgehog (Shh) signaling

(Tran, et al., 2008). The mutation spectrum of CEP290 is even more striking as pathogenic variants have been associated with a diverse array of ciliopathies without any obvious genotype- phenotype correlations (reviewed in Coppieters, et al., 2010). Functionally, the CEP290 protein localizes to both the cilia transition zone (Craige, et al., 2010) and to centriolar satellites (Kim, et al., 2008a; Stowe, et al., 2012), where it interacts with a number of other ciliary proteins required

for cilia formation (Stowe, et al., 2012). Knockdown of CEP290 in cultured mammalian cells disrupts satellite localization, preventing assembly of a functional cilium (Stowe, et al., 2012).

Similar disruption of centriolar satellites by morpholino-mediated knockdown of pericentrin

(pcm1) in zebrafish yielded inverted heart looping in nearly 50% of morphant embryos (Stowe, et al., 2012). These results suggest pathogenic potential for CEP290 mutations via loss of centrosomal protein targeting and ciliary assembly. While CEP290 mutations have not yet been reported as causes of human heterotaxy, at least one patient with SI has been identified to carry a

CEP290 mutation (Brancati, et al., 2007).

Two other ciliary genes, TTBK2 and CFAP126, were identified in CNVs in the heterotaxy cohort. The first, TTBK2, encodes a serine/threonine kinase essential for IFT protein recruitment. Its localization to centrioles is an initiating step of ciliogenesis and is required to promote elongation of the ciliary axoneme (Goetz, et al., 2012). Like TTC21B and CEP290, mutations in TTBK2 prevent formation of a functional cilium and are a recognized cause of disease in both humans (spinocerebellar ataxia 11) and bartleby (bby) mutant mice. TTBK2 mutations have yet to be reported among patients with heterotaxy; however, mice homozygous for bby mutations exhibit randomized heart looping indicative of impaired L-R patterning, in addition to other defects stemming from disrupted ciliogenesis (holoprosencephaly, limb defects, twisted body axis). Comparatively less is known about CFAP126, a newly characterized protein important for basal body docking and cilia positioning (Gegg, et al., 2014). Nevertheless, expression has been noted at the mouse embryonic node as early as E7.75, placing CFAP126 at the correct location and time for impacting laterality (Lange, et al., 2012).

Alongside these ciliary genes, two rare CNVs encompassing members of the TGF superfamily were identified. The first, a duplication involving the Nodal signaling modulator,

NOMO3, was detected in a male patient with sporadic heterotaxy (Patient 19). NOMO3 exists as one of three highly similar genes on chromosome 16p12-p13 and is thought to act as a direct

Nodal signaling antagonist, acting independently of other Nodal modulating pathways (Haffner, et al., 2004). The second CNV, a duplication identified in a male patient with familial heterotaxy

(Patient 40), encompassed LTBP1, a TGFbinding protein with roles in TGF1 assembly, secretion, and targeting. Deletions involving LTBP1 have been previously associated with outflow tract (OFT) defects (Todorovic, et al., 2007), but have not yet been reported among patients with heterotaxy. Intriguingly, experiments in Xenopus have identified LTBP1 as a potent activator of Activin and Nodal signaling, indicating potential for involvement in L-R relevant pathways (Altmann, et al., 2002).

A ~200kb deletion involving the GJB6 gene, which encodes the gap junction protein connexin 30, was identified in a prenatally diagnosed female patient with hydrocephalus, meningomyelocoele, and complex CHD (Patient 35). While GJB6 has not yet been linked to heterotaxy, mutations in GJA1 (connexin 43) have been previously reported in six patients with heart malformations and laterality defects (Britz-Cunningham, et al., 1995). These results have subsequently been challenged as follow-up screenings failed to identify similar mutations in other heterotaxy populations (Casey and Ballabio, 1995; Gebbia, et al., 1996; Debrus, et al.,

1997; Penman Splitt, et al., 1997). Nevertheless, as gap junction-mediated transport has been proposed as a key component of the “ion-flux” model of early L-R patterning in chick, frog, and zebrafish (Levin and Mercola, 1998; Levin and Mercola, 1999; Vandenberg, et al., 2014), and in determination of L-R neuronal asymmetry in Caenorhabditis elegans (Chuang, et al., 2007), involvement of gap junctions in human laterality programs cannot be entirely ruled out.

Rare CNVs involving four distinct microRNAs (miR-148a, miR-455, miR-484, and miR-

2117) were also identified in the heterotaxy cohort. Considering known functions in the context of L-R patterning, miR-418a is the most notable as it is a direct inhibitor of both ACVR1, an activin receptor (Song, et al., 2012) and ROCK1, a rho-associated kinase (Zhang, et al., 2012; Li, et al., 2013). Although neither protein has been directly associated with laterality, family members of both (ACVR2B, ROCK2) are known genetic causes of heterotaxy in human patients

(Kosaki, et al., 1999b; Fakhro, et al., 2011).

A total of ten rare CNVs were identified at loci previously associated with CHD (Supp.

Table 2.10.4). To the best of our knowledge, only one of these, a large 1.7-2.6Mb 2q13 duplication detected by oligoarray in a male patient with sporadic heterotaxy and complex CHD

(Patient 29), affects a locus previously associated with heterotaxy (Rudd, et al., 2009). The duplicated region encompassed 10 genes (Supp. Table 2.10.2) and was identical in gene content to a 1.62Mb deletion detected in a patient with a similar laterality phenotype (Rudd, et al., 2009).

Genomic imbalances spanning ~1.7Mb and encompassing the same 10 genes have been described in a number of other patients CHD but who are otherwise lacking overt situs abnormalities (Rudd, et al., 2009; Cooper, et al., 2011; Soemedi, et al., 2012; Yu, et al., 2012;

Russell, et al., 2014). CHD is more common to deletion than duplication carriers; however, in many cases, results from cardiac evaluations have not been available (reviewed in Yu, et al.,

2012). No specific microduplication/deletion syndrome has been defined in the region; however, brain abnormalities were suggestive of Joubert syndrome in one individual (Yu, et al., 2012).

The CNV interval in this patient did not encompass the Joubert genes, NPHP1 and AH11, which were fully sequenced and found to be normal (Yu, et al., 2012). Recent knockdown studies in zebrafish have identified FBLN7 and TMEM87B as the genes in the genomic interval most likely

69 to be causative for CHD; however, their exact roles in cardiac development have yet to be determined (Russell, et al., 2014). In our patient, a second, larger CNV was also identified upstream of this region and included an additional 18 genes of uncertain pathogenic significance.

Predicted breakpoints of the two intervals were non-overlapping but were in immediate proximity, suggesting the possibility of a single duplication event.

2.5.2. PFKP as a novel cause of heterotaxy

PFKP, encoding the platelet isoform of the glycolytic enzyme, phosphofructokinase-1

(PFK-1) was among the top heterotaxy candidates genes identified in the cohort. Using established morphlino-based methods for candidate gene screening (Fakhro, et al., 2011; Cast, et al., 2012; Boskovski, et al., 2013), we have identified pfkp as a novel regulator of laterality in

Xenopus laevis.

PFK-1 is the rate-limiting enzyme of glycolysis, catalyzing the irreversible conversion of fructose 6-phosphate to fructose 1,6-bisphosphate. In both human and frog, PFK-1 exists as three isoforms, each encoded by a unique locus and named in accordance with adult expression

(muscle, M; liver, L; and platelet/fibroblast, P). Recessive loss of function mutations in PFKM result in diabetes as well as glycogen storage disease type VII (Tarui’s disease) (Ristow, et al.,

1997). Our interest in PFKP was therefore additionally piqued by an unexplained, but long- recognized, increased risk for heterotaxy among the children of diabetic mothers (Morishima, et al., 1996; Slavotinek, et al., 1996; Splitt, et al., 1999; Loffredo, et al., 2001; Martinez-Frias,

2001; Lisowski, et al., 2010; Lopez, et al., 2015).

Our lab (Figure 2.9.2, panel b) and others (Davidson, et al., 1983; Raddatz and Lovtrup-

Rein, 1986; Dworkin and Dworkin-Rastl, 1991) have shown that PFK-1 is expressed throughout

early Xenopus development. Expression is most readily detected in the animal pole and marginal zone during gastrulation, and at high levels in the neural tube, neural crest, and adjacent tissue during neurulation (Pegoraro, et al., 2013). Overall levels of expression increase as development proceeds, particularly in dorsal tissues (Pegoraro, et al., 2013). At least in humans, there does not appear to be a specific “fetal” isoform as all three types are uniformly expressed in early embryonic stages: it is only as organ systems begin to develop that isoform-specific expression patterns begin to emerge (Kahn, et al., 1980; Davidson, et al., 1983). Intriguingly PFKP is the sole isoform identified at significant levels in fetal heart, which contrasts sharply with a preponderance of PFKM in adult cardiac tissue (de Faria, et al., 1978; Kahn, et al., 1980). The significance of this finding, particularly with respect to CHD and heterotaxy, is presently uncertain. Because our morpholinos were designed to explicitly target the 5’ untranslated region of PFKP and not the M and L subtypes (which are encoded by different genes), it also remains to be seen whether embryonic knockdowns of these isoforms similarly disrupt L-R patterning

In Xenopus, catalytic activity of PFK-1 isoforms (Raddatz and Lovtrup-Rein, 1986) and overall rates of glycolysis (Dworkin and Dworkin-Rastl, 1991) are very low until gastrulation

(Dworkin and Dworkin-Rastl, 1991). As L-R patterning mechanisms may operate during early cleavage stages (Levin, et al., 2002; Levin, 2004; Fukumoto, et al., 2005b; Fukumoto, et al.,

2005a; Adams, et al., 2006; Aw, et al., 2008; Morokuma, et al., 2008; Aw, et al., 2010;

Vandenberg, et al., 2013), roles for PFKP in L-R patterning could manifest during pre-gastrula stages. Possible non-glycolytic functions for PFKP must therefore also be considered. It is potentially noteworthy that a non-glycolytic role in early dorsal patterning has recently been demonstrated for PFKFB4, one of five known isoforms of phosphofructokinase-2 (PFK-2)

(Pegoraro, et al., 2015).

How might PFKP regulate L-R patterning independently of glycolysis? Isoforms of PFK-

1 are known to bind the a-subunit of H+-V-ATPase (Su, et al., 2003; Su, et al., 2008), a plasma membrane and vacuolar proton pump whose inhibition in pre-gastrula stages results in significant defects in a number of L-R patterning processes including ion-flux (Xenopus/chick),

LRO and cilia development (zebrafish), and Nodal signaling (chick/zebrafish) (Adams, et al.,

2006; Gokey, et al., 2015). Together with observations from yeast that PFK-1 is required to maintain activity of the H+V-ATPase proton pump and that even a catalytically inactive PFK-1 is capable of completing this function (Su, et al., 2008), a reasonable hypothesis is that PFKP knockdown may impact L-R patterning through loss of a functionally critical interaction with the

H+-V-ATPase a-subunit. As recent zebrafish studies indicate a primary role for H+-V-ATPase in

KV formation (Gokey, et al., 2015), it is possible that PFKP may impact L-R patterning in a similar fashion. Interestingly, a rare duplication involving the gene ATP6V1G1, which encodes the G-subunit of H+-V-ATPAase, was also identified in a male heterotaxy patient in our cohort

(Patient 41). Like the a-subunit, the G-subunit contributes to the peripheral stator stalk, the non- rotating connection between the V1 and Vo halves of the ATPase (Beyenbach and Wieczorek,

2006). Two of three dimers composed of E and G-subunits attach to the a-subunit at its amino terminal domain, a site thought to be of importance for a-subunit function (reviewed in Holliday,

2014). Our initial attempts at dose-synergy experiments using sub-threshold dosages of pfkp and

H+-V-ATPase a-subunit (atp6voa1) morpholinos were inconclusive (data not shown); however, future experimentation should help to definitively establish whether the PFKP / H+-V-ATPase interaction is truly functionally relevant to laterality.

2.5.3. STUDY LIMITATIONS

Study design was subject to a number of limitations common to investigations of this nature. As blood samples from parents of all study participants were not available, comprehensive determinations of de novo inheritance were not possible. Similarly, while a few instances of copy number mosaicism were identified, direct comparisons between mosaic and non-mosaic CNV carriers were prevented by the rarity of individual CNVs. Cut-offs for CNV calling were also set at a minimum coverage of 3 markers/probes per CNV, restricting downstream genetic and functional analyses to variants exceeding these minimum size thresholds. While more difficult to detect reliably by microarray approaches, small exonic CNVs of 1-30kb have been suggested to contribute to susceptibility of some genetic diseases (Poultney, et al., 2013). Finally, analyses were restricted to CNVs involving coding regions of at least one gene, ignoring variants solely impacting intronic or regulatory sequences. Genetic variation in non-coding regions, while typically of less obvious functional significance, has been linked to genetic disease (Maurano, et al., 2012; Ward and Kellis, 2012; Khurana, et al., 2013) but would not have been considered in our study.

Our analyses of PFKP as a L-R patterning candidate were also limited by a number of factors. First, we were unable to definitively verify isoform-specific knockdown by Western blotting as the M and L isoforms are of similar size and commercially available antibodies demonstrate high likelihood for subtype cross-reactivity (data not shown). Nevertheless, confidence in the specificity of TB knockdown was provided by replication of the TB morphant phenotype in SB morphants and demonstration that sub-threshold dosages of both morpholinos synergized to produce organ situs defects when used in combination, but not in isolation.

Second, while we have demonstrated disruption of normally right-sided coco expression in pfkp

73 morphants, demonstrations of effect on other asymmetric markers (xnr-1/pitx2) will be helpful in definitively confirming abrogation of left-sided Nodal signaling. Finally, our work on this project pre-dated emergence of CRISPR/Cas9 methods as an efficient means for achieving gene-specific knockdown (reviewed in Sander and Joung, 2014). While the pseudotetraploid genome of

Xenopus laevis complicates use of these technologies in our existing colony, they can more easily be applied to diploid Xenopus tropicalis (Blitz, et al., 2013; Nakayama, et al., 2013). As recent zebrafish studies have highlighted the potential for discrepancies between morphant and mutant phenotypes (Kok, et al., 2015; Stainier, et al., 2015), future work should endeavor to confirm the effects of pfkp knockdown on L-R patterning using CRISPR/Cas9 or equivalent genetic knockdown methods.

2.6. FUTURE DIRECTIONS

The utility of array-based CNV analyses for the study of heterotaxy has been now been definitely demonstrated, with present and past reports identifying a number of novel heterotaxy genes and candidate loci for future study (Fakhro, et al., 2011; Rigler, et al., 2014; this study).

What these investigations share is an understanding that the genetics of heterotaxy cannot be fully explained by simple Mendelian inheritance alone and that unbiased genome-wide approaches are required to delineate previously unconsidered genes and pathways. The introduction of similarly unbiased next-generation sequencing (NGS) technologies has already begun to revolutionize the study of complex genetic disease, (Casals, et al., 2012; Handel, et al.,

2013; Kilpinen and Barrett, 2013), identifying novel genetic causes for a growing list of developmentally and genetically heterogeneous disorders, including heterotaxy (Tariq, et al.,

2011). Nevertheless, despite advantages in speed, cost, and resolution over traditional microarray

techniques, NGS methods for the study of copy number variation remains hampered by a lack of comparably robust statistical approaches to CNV identification (Wang, et al., 2014). While this is a serious limitation, continued refinement of NGS technologies and active development of novel statistical methods (Duan, et al., 2013; Zhao, et al., 2013; Tan, et al., 2014; Chen, et al.,

2015) indicate that these technical obstacles may soon be overcome, clearing the way for more widespread application of NGS to the study of copy number variation.

2.7. ACKNOWLEDGEMENTS:

We thank the patients and families for their participation. This work was supported by funding from the Burroughs Welcome Fund (#1008496) and March of Dimes Research

Foundation (#1FY10-401) (to S. Ware.).

2.8. TABLES

Table 2.8.1. Heterotaxy cohort demographics.

Gender Inheritance Racial and Ethnic Categories Totals (%)

Females Males Familial Sporadic Unknown

Arabic 2 (0.9) 2 0 0 2 0

Asian 2 (0.9) 0 2 0 2 0

Black or African American 13 (5.8) 8 5 1 9 3

Caucasian 90 (40.0) 33 57 22 54 14

Hispanic / Latino 68 (30.2) 21 47 8 48 12

Mixed 8 (3.6) 3 5 3 5 0

Unknown 42 (18.7) 19 23 12 9 21

Totals (%) 225 (100.0) 86 (38.2) 139 (61.8) 46 (20.4) 129 (57.3) 50 (22.2)

Percentages are in parentheses and relative to the 225 patient total.

Table 2.8.2. Pathogenic chromosome abnormalities detected in the 225 patient heterotaxy cohort.

Cytogenetic findings (breakpoints)2 Patient Gender Type Platform1 Inheritance / Phenotype Illumina SNP array Agilent oligo array 1 M gain oligo na arr 9q33.2-q33.3(125204518-140138901)x3 sporadic / l-TGA, PA, ASD, VSD unbalanced translocation / arr 5q15q35.3(94068223-176886171)x3 2 M oligo na familial / d-TGA, CAVC, VSD, ASD, asplenia complex arr 9p24.1p23(8442856-12223883)x1 rearrangement sporadic /situs ambiguus, abd. SI, VSD, DORV, PS, PA, SVC abnls., asplenia, renal abnls., pulmonary isomerism, heart malrotation, 3 M loss 1M, oligo arr 22q11.1q11.21(15815765-16422571)x1 arr 22q11.1q11.21(15815765-16422571)x1 hypoplastic pulmonary artery, central gallbladder, pancreas rotated to Rt., stomach rotated to Lt. , hypoplastic multicystic Rt. kidney, undescended Lt. testicle, absent Rt. testicle. 2p12p11.2(82350453-84912738)x3, 11p14.2q13.3(26301682-70604313)x2 hmz familial / oligohydramnios, hypoplastic lungs; 4 M gain; UPD 1M na UPD around centromere from11p14.2- ASD, PAPVR, DORV, CAVC, dextrocardia 11q13.3 (26301682-70604313) 7p22.3p21.3(34332-12922987)x1-2 7p21.3p15.1(12925517-28332937)x1-2 hmz unknown / SI; PDA; Kartagener’s: immotile cilia mosaic gain 7p15.1p12.3(28,333,319-48,346,413)x1-2 arr 7p22.3p12.3(136551-48338627)x1 5 F 1M, oligo by nasal biopsy; consanguinity (parents 1st and loss 7p12.1(48,346,704-48,898,843)x2-3 arr 7p11.2(53925715-56351979)x3 cousins) 7p12.3p12.1(48900806-51511550)x2 7p12.1p11.2(51513509-56362278)x2-3 arr 2p25.3(3,274,169-3,431,910)x1 unknown / Rt. aortic arch, VSD; abd. SI; gut 6 F loss oligo na arr 22q11.2(17020267-19761174)x1 malrotation sporadic / DORV, ASD, VSD, PS, polysplenia, mitral atresia, duplicated IVC, midline pancreas, hypoplastic scrotum, microcephaly (FOC 45cm), 7 M loss 1M, oligo arr 8p23.3p21.2(146703-25298793)x1 arr 8p23.3p21.2(151,470-25,293,991)x1 bilat. medial ulegyria of frontal hemispheres, organomegaly (liver, spleen, kidneys >95% for age) mosaic sporadic / dextrocardia, SVC abnls., hypoplastic 8 M 1M, oligo arr 21q11.2q22.3(13989691-46942324)x1-2 arr 21q11.2q22.3(13971999-46914745)x1 monosomy 21 Rt. thumb, chordee arr 2q35(216859560-217354800)x3 arr 2q35(216852776-217347523)x3 complex sporadic / d-TGA, HLHS, VSD, ASD, mitral 9 M 1M, oligo arr 18p11.32p11.31(99027-6681691)x1 arr 18p11.32p11.31(121700-6596548)x1 rearrangement atresia, hypoplastic LV, subvalvar and valvar PS arr 18p11.31p11.21(6683107-15400816)x3 arr 18p11.31p11.21(6684807-14969496)x3 arr 2q37.3(237239911-242692820)x3, unbalanced familial / dextrocardia with Rt. aortic arch, atrial 10 F 370k arr 10q26.13q26.3(125436928- na translocation inversion, CAVC, SVC abnls., abd. SI, asplenia 135320785)x1 mosaic (2)x2 hmz mos, sporadic / unbalanced AVC, single outlet RV, 11 M unbalanced 1M, oligo 7q22.1q36.3(98613389-158815293)x2-3, na TAPVR; abd. SI, asplenia translocation 10q22.1q26.3(73126274-135374311)x1-2 12 M mosaic gain 1M, oligo arr 5q11.2q35.3(51392113-180837635)x2-3 entire 5q (88Mb) sporadic / CHD sporadic / d-TGA, VSD, ASD, DORV, mod. dil. 13 F loss 1M arr 1p34.3p34.1(36507566-45013872)x2-3 na RV/RA, mild RV hypertrophy, large tortuous PDA, excessive aortopulmonary collaterals, DD

(MRI: 2 small sites of leukomalacia in white matter of frontal lobes) 2For the purposes of this and all subsequent tables, array-CGH is listed as oligo array, while SNP-arrays are designated as 370 or 1M depending on number of included markers (370,000 vs. 1,000,000). 1Linear positions according to Build 36, hg18. Standard cytogenetic nomenclature is used. Abbreviations: abd. = abdominal; abnl/abnls. = abnormal/abnormalities; AS = aortic stenosis; ASD = atrial septal defect; AVC = atrioventricular canal; AVVR = atrioventricular valve regurgitation bilat. = bilateral; CAVC = complete atrioventricular canal; CNV = copy number variant; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DD = developmental delay; dil. = dilated; DILV = double inlet right ventricle; DORV = double outlet right ventricle; HLHS = hypoplastic left heart syndrome; IVC = inferior vena cava; l-TGA = levo-transposition of the great arteries; Lt. = left; LV = left ventricle; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PDA = patent ductus arteriosus; PS = pulmonic stenosis, Rt. = right; RA = right atrium; RV = right ventricle; SI = situs inversus; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; VSD = ventricular septal defect

Table 2.8.3. Rare CNVs encompassing known heterotaxy genes.

Size Size Agilent # # Affected Patient Gender Type Locus Platform Illumina (min-max) Markers Probes Inheritance / Phenotype Reference Genes (bp) (bp) (Illumina) (Agilent)

Gebbia, et al., 1997; Megarbane, et al., 2000; Ware, et al., 2004; sporadic / dextrocardia, single Fritz, et al., 2005; Tzschach, et ventricle, CAVC, AVVR, PA, al., 2006; Chhin, et al., 2007; De 14 M loss Xq26.2 1M, oligo 2876 3312 2 3 ZIC3 PS, asplenia, mod. enlarged Lt. Luca, et al., 2010; Wessels, et al., hepatic lobe (Deceased) 2010; Chung, et al., 2011; D'Alessandro, et al., 2011; Ma, et al., 2012; Cowan, et al., 2014 ROCK2, E2F6, not not sporadic / d-TGA, CoA, double 15 M gain 2p25.1 oligo 553886-685571 13 GREB1, Fakhro, et al., 2011 analyzed analyzed chambered RV/DORV, VSD NTSR2, LPIN1 familial (Sister with ASD) / abd SI, PAPVR, abnl. SVC, RBMS3, 1457325- abnl. IVC, VSD, ASD, 16 F loss 3p24.1 1M, oligo 1491644 617 76 TGFBR2, Fakhro, et al., 2011 1606574 polysplenia, slightly upslanting GADL1 palpebral fissures, long upper lip Abbreviations: Abd. = abdominal; abnl. = abnormal; ASD = atrial septal defect; AVVR = atrioventricular valve regurgitation CAVC = complete atrioventricular canal; CNV = copy number variant; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DORV = double outlet right ventricle; IVC = inferior vena cava; mod. = moderately; Lt. = left; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PS = pulmonary stenosis, RV = right ventricle; SI = situs inversus; VSD = ventricular septal defect

2.9. FIGURES

Figure 2.9.1. Overview of CNV findings. CNVs were designated as “common” or “rare” based on their presence in clinical and online databases of genomic variation (see Chapter 2.3.3). CNVs denoted as having “clear pathogenicity” included a diverse array of large/complex chromosomal abnormalities (Table 2.8.2), as well as CNVs containing genes previously associated with heterotaxy (Table 2.8.3). “CNVs of interest” represent CNVs containing genes of potential interest for candidate gene studies.

Figure 2.9.2. pfkp but not pitrm1 Xenopus laevis morphants develop organ situs defects by st. 47. (a) Genes in CNV interval. Source: USCS genome browser (https://genome.ucsc.edu/) (b) RT-PCR results demonstrate that pfkp is expressed throughout all critical L-R patterning stages. ODC = ornithine decarboxylase (c) pfkp knockdown in 2-cell embryos causes heterotaxy in Xenopus. Situs defects in st. 47 morphants are shown relative to zic3 control morphants and uninjected controls. (d) Confirmation of exon 8 deletion in pfkp SB morphants by RT-PCR targeting the exon 8-intron 8 splice junction. Upper schematic: Small horizontal arrows denote primer annealing sites. The horizontal line without an arrowhead denotes the splice block morpholino target site. The severely truncated exon 8 (see text) is represented by cross-hatching. Lower panel (RT-PCR): Large arrowhead denotes the pfkp deletion fragment. SB = pfkp splice block

Figure 2.9.3. Synergy and rescue experiments support MO specificity. (a) Sub-threshold dosages of pfkp TB and SB MOs yielded a higher proportion of embryos with situs defects when injected together than when injected alone. (b) Partial rescue of organ situs in SB morphants co- injected with WT human PFKP mRNA. SB = splice block, TB = translation block

Figure 2.9.4. pfkp knockdown disrupts right-sided coco bias in st. 20-21 embryos. (a) Representative images of coco expression at the gastrocoel roof plate (GRP) in st. 20-21 embryos. (b) Schematic depicting right-sided coco bias during post-flow stages. Reduced coco expression on the left-side permits initiation of the Nodal signaling cascade by alleviating inhibition of Xenopus Nodal related-1 (Xnr-1). Downstream transcription factors (Pitx2) important for initiating asymmetric organ morphogenesis are activated. High levels of right-sided coco expression repress similar activation of Nodal signaling on the right side. (c) Relative to control embryos, significantly fewer pfkp morphants exhibited right-sided bias of coco expression (p = 0.0058, two tailed Fisher’s Exact test).

2.10. SUPPLEMENTARY MATERIALS

Supp. Figure 2.10.1. Gene ontologies for genes identified in 35 CNVs of interest. All GO enrichment analyses were completed using PANTHER v. 10.0 (http://pantherdb.org/).

Supp. Table 2.10.1. Phenotypic summary of the 225 patient heterotaxy cohort.

Patients with pathogenic or Heterotaxy likely pathogenic Patients with Cohort CNVs common CNVs % % % Overall Phenotype: n n n p-value1 (n/225) (n/49) (n/176) SI totalis 18 8.00 4 8.16 14 7.95 1.0000 (NS) Heterotaxy 153 68.00 39 79.59 114 64.77 0.0572 (NS) Isolated CHD 43 19.11 5 10.20 38 21.59 0.0990 (NS) VACTERL-like 4 1.78 0 0.00 4 2.27 0.5790 (NS) Normal, Familial disease 1 0.44 0 0.00 1 0.57 1.0000 (NS) No phenotype details 6 2.67 1 2.04 5 2.84 1.0000 (NS) Abdomen/gastrointestinal:

Abdominal SI 82 36.44 23 46.94 59 33.52 0.0948 (NS) Bile duct hypoplasia/biliary atresia 3 1.33 0 0.00 3 1.70 1.0000 (NS) Gallbladder abnormalities 6 2.67 1 2.04 5 2.84 1.0000 (NS) Malrotation of the gut 48 21.33 9 18.37 39 22.16 0.6944 (NS) Asplenia 63 28.00 13 26.53 50 28.41 0.8588 (NS) Polysplenia 22 9.78 4 8.16 18 10.23 0.7911 (NS) Multilobated spleen 1 0.44 0 0.00 1 0.57 1.0000 (NS) HJ bodies 5 2.22 2 4.08 3 1.70 0.2989 (NS) Imperforate anus 3 1.33 0 0.00 3 1.70 1.0000 (NS) Renal anomalies 11 4.89 2 4.08 9 5.11 1.0000 (NS) Liver anomalies 35 15.56 5 10.20 30 17.05 0.2752 (NS) Skeletal/limbs

Vertebral anomalies 10 4.44 2 4.08 8 4.55 1.0000 (NS) Rib anomalies 4 1.78 1 2.04 3 1.70 1.0000 (NS) Scoliosis 3 1.33 1 2.04 2 1.14 0.5232 (NS) Limb defects 6 2.67 1 2.04 5 2.84 1.0000 (NS) Cardiac position:

Levocardia 20 8.89 4 8.16 16 9.09 1.0000 (NS) Mesocardia 5 2.22 1 2.04 4 2.27 1.0000 (NS) Dextrocardia 13 5.78 2 4.08 11 6.25 0.7388 (NS) Atria/Ventricles:

Common atrium 16 7.11 1 2.04 15 8.52 0.2048 (NS) Atrial isomerism 10 4.44 0 0.00 10 5.68 0.1232 (NS) ASD 76 33.78 20 40.82 56 31.82 0.3054 (NS) AV canal 55 24.44 11 22.45 44 25.00 0.8513 (NS) Single ventricle / HLHS 37 16.44 10 20.41 27 15.34 0.3902 (NS) VSD 98 43.56 20 40.82 78 44.32 0.7453 (NS) Vessels:

SVC abnormality 49 21.78 14 28.57 35 19.89 0.2396 (NS) IVC abnormality 49 21.78 7 14.29 42 23.86 0.1745 (NS) TAPVR 32 14.22 7 14.29 25 14.20 1.0000 (NS) PAPVR 16 7.11 6 12.24 10 5.68 0.1230 (NS) Inflow/Outflow:

Aortic arch abnormalities 57 25.33 15 30.61 42 23.86 0.3560 (NS) d-TGA 81 36.00 9 18.37 72 40.91 0.0040 (S) l-TGA 31 13.78 8 16.33 31 17.61 1.0000 (NS) DILV 14 6.22 1 2.04 13 7.39 0.3129 (NS) DORV 65 28.89 19 38.78 46 26.14 0.1081 (NS) PA 49 21.78 12 24.49 37 21.02 0.6956 (NS) PS 58 25.78 11 22.45 47 26.70 0.5859 (NS) AS 4 1.78 2 4.08 2 1.14 0.2074 (NS) CoA 22 9.78 3 6.12 19 10.80 0.4233 (NS) BAV 2 0.89 1 2.04 1 0.57 0.3889 (NS) Other:

Arrhythmia 16 7.11 2 4.08 14 7.95 0.5325 (NS) 1 p-values for statistical significance (p<0.05) represent comparisons between patients with CNVs of clear or potential pathogenicity and patients with common CNVs by two-tailed Fischer’s Exact tests using Graphpad statistical software (http://www.graphpad.com/quickcalcs/). NS = non- significant, S = significant.

Abbreviations: AS = aortic stenosis; ASD = atrial septal defect; CAVC = complete atrioventricular canal; BAV = bicuspid aortic valve; CHD = congenital heart disease; CNV = copy number variant; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DILV = double inlet right ventricle; DORV = double outlet right ventricle; HJ bodies = Howell-Jolly bodies; IVC = inferior vena cava; l-TGA = levo- transposition of the great arteries; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PS = pulmonic stenosis, including sub, valvar, and supravalvar but not branch PS; SI = situs inversus; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; VACTERL = vertebral, anal, cardiac, tracheo-esophageal, renal, radial, limb; VSD = ventricular septal defect

Supp. Table 2.10.2. Rare coding region CNVs detected in the 225 patient heterotaxy cohort.

Size Size Start Stop Min Start Max Start Min Stop Max Stop Agilent # Markers # Probes Patient Gender Type Locus Platform Illumina Affected Genes Inheritance / Phenotype (Illumina) (Illumina) (Agilent) (Agilent) (Agilent) (Agilent) (min-max) (Illumina) (Agilent) (bp) (bp)

sporadic / situs ambiguus, abd. SI, gut malrotation, single ventricle, PS, MGA, 370k, 2517224- 17 M gain 2q32.1 183850093 186628963 183848222 183820875 186365446 186401261 2778870 246 50 ZNF804A DILV, TAPVR, PAPVR, oligo 2580386 bilat. SVC, VSD, PDA, asplenia, midline liver, cleft lip/palate sporadic / levocardia, dextrocardia, abd. SI, gut n/a, n/a, n/a, n/a, n/a, n/a, n/a, 18 F gain 15q13.3 1M, oligo 579670 174 n/a CHRNA7 malrotation, HLHS, validated validated validated validated validated validated validated CAVC, PS, DORV, TAPVR, abnl. IVC, ASD C16orf45, KIAA0430, NDE1, MIR484, sporadic / levocardia, situs gain 16p13.11 not not not 2282653- not 19 M oligo 15429214 15062247 17711867 17962996 326 MYH11, FOPNL, ambiguus, abd. SI, PS, l- -p12.3 analyzed analyzed analyzed 2900749 analyzed ABCC1, ABCC6, TGA NOMO3, XYLT1 TXNIP, POLR3GL, ANKRD34A, LIX1L, RBM8A, GNRHR2, not not not 300679- not PEX11B, ITGA10, 19 gain 1q21.1 oligo 144151335 144124804 144452014 145085612 17 analyzed analyzed analyzed 960808 analyzed ANKRD35, PIAS3, NUDT17, POLR3C, RNF115, CD160, PDZK1 sporadic / dextrocardia, abd. SI, PS, DORV, TAPVR, abnl. SVC, ASD, not not not 14097- not asplenia, atrial SI, mitral 20 M gain 7q21.11 oligo 83513348 83502873 83527445 83527519 4 SEMA3A analyzed analyzed analyzed 24646 analyzed hypoplasia with mitral stenosis, subvalvar PS, possible midline pancreas, sacral dimple familial (Aunt with HLHS) / levocardia, abd. SI, CAVC, PA, Rt. aortic arch, d-TGA, abnl. SVC, VSD, asplenia, n/a, n/a, n/a, n/a, hydrocephalus, 21 M loss 4q32.2 1M, oligo 162862230 163211975 350000 331000 101 n/a FSTL5 validated validated validated validated coagulation defect, increased fibrin split products, decreased platelets, PIT, bilat., renal enlargement, mild pyeloectasia, maternal diabetes ZNF480, ZNF610, sporadic / dextrocardia, 120782- ZNF610, ZNF880, PA, l-TGA, VSD, ASD, 22 F loss 19q13.33 1M, oligo 57510893 57672568 57532622 57505693 57653404 57668223 162000 119 4 162530 ZNF528, ZNF534 arrhythmia (Pacemaker), ZNF578 scoliosis familail (pat. 1st cousin with septal 10071- 23 F loss 12q21.32 oligo not called not called 87010586 87010537 87020657 87020686 not called not called 7 CEP290 defect) / gut malrotation, 10149 d-TGA, VSD, ASD, d- looped ventricles, IAA,

subvalvular AS, RA/RV dilation, hypoplastic spleen, sacral dimple not not not 618317- not 24 na gain 12q24.32 oligo 124595123 124554733 125213440 125263456 15 TMEM132B na analyzed analyzed analyzed 708723 analyzed familial (1st cousin with renal anomaly) / SI totalis, TAPVR, arch abnls., Rt. aortic arch., dextrocardia not not not 208827- not 25 M loss 6q26 oligo 162394936 162357384 162603763 162784160 14 PARK2 with atrioventricular and analyzed analyzed analyzed 426776 analyzed ventricularterial concordance, hemivertebrae, microtia (Deceased). familial (male sib expired w/same defects.) / situs ambiguus, arch abnls, d- TGA, DORV, VSD, ASD, 261043- anterior & rightward aorta, 26 M loss 8q22.2 1M, oligo 100293643 100580386 100301924 100291476 100562967 100563091 286743 32 38 VPS13B 271615 severe valvar/subvalvar PS, small MPA, atretic LPA, RAA, Rt. ear lobe "pushed up", foot positional defects familial / situs ambiguus, ASD, AVVR, single ventricle, gut malrotation, not not not 67216- Not liver abnls, dysplastic 27 M loss 7p21.1 oligo 15550929 15550789 15618145 15665559 17 MEOX2, AGMO analyzed analyzed analyzed 114770 analyzed single AV valve, TGA with aorta anterior and pulmonary artery posterior, enlarged RA sporadic / situs ambiguus, dextrocardia, abd. SI, single ventricle, absence of pulmonary trunk, bilat. TRNAE-UUC, SVC/IVC, arch abnls., 13q14.11 n/a, n/a, n/a, n/a, n/a, 28 F loss 1M, oligo 44150171 44780162 629991 14 n/a NUFIP1, GPALPP1, TAPVR, ASD, bilat. -q14.12 validated validated validated validated validated GTF2F2, KCTD4 trilobed lungs, asplenia, bilat. mild hydronephrosis, mother on Tegretol for seizures during pregnancy (Deceased) n/a, n/a, n/a, n/a, n/a, 28 gain 6q11.1 1M, oligo 61942860 62980807 1037947 223 n/a KHDRBS2 validated validated validated validated validated ST6GAL2, RGPD4- AS1, RGPD4, SLC5A7, SULT1C3, sporadic / dextrocardia, SULT1C2, SULT1C4, ASD, PA, atrial inversion, 2q12.2- not not not 3683400- not GCC2, LIMS1, 29 M gain oligo 106499382 106350195 110182782 110183109 208 common AV valve, single q13 analyzed analyzed analyzed 3832914 analyzed RANBP2, CCDC138, RV; asplenia, multiple EDAR, SH3RF3-AS1 sinus infections, ADHD , SH3RF3, SEPT10, SOWAHC, LIMS3, LOC440895 BUB1, ACOXL, BCL2L11, MIR4435- not not not 1695755- not 2HG, ANAPC1, 29 gain 2q13 oligo 111123310 110340992 112819065 112926881 67 analyzed analyzed analyzed 2585889 analyzed MERTK, TMEM87B, FBLN7, ZC3H8, ZC3H6 sporadic / dextrocardia, abd. SI, d-TGA, VSD, n/a, n/a, n/a, n/a, n/a, n/a, multicystic dysplastic Lt. 30 F gain 9p23 1M, oligo 307240 508746 210506 196 DOCK8, KANK1 validated validated validated validated validated validated kidney, 2-vessel umbilical cord with umbilical artery directly from aorta, caudal

regression/sacral agenesis, lower limb malrotation, bilat. clubbed feet (Deceased) sporadic / HLHS, CoA, arch abnls., d-TGA, n/a, n/a, n/a, n/a, n/a, 31 F loss 7q22.1 oligo not called not called not called 98000 not called NPTX2 DORV, ASD, mitral validated validated validated validated validated atresia with severely hypoplastic LV not not not 111657- not sporadic / SI totalis, 32 F gain 15q15.2 oligo 40951832 40937795 41063489 41063520 59 TTBK2, UBR1 analyzed analyzed analyzed 125725 analyzed ?Kartagener syndrome sporadic / situs ambiguus, HLHS variant, CAVC, ASD, aortic atresia, hypoplastic LV, mod. RV dilatation, interrupted midline IVC with hemiazygous cont. to Lt. n/a, n/a, n/a, n/a, n/a, 33 M loss 16p13.12 oligo not called not called not called 287000 not called SNX29 SVC, absent Rt. SVC, Lt. validated validated validated validated validated SVC to dilated coronary sinus, pulmonary vein entrance into LA distorted by large coronary sinus, pulmonary artery dilatation, ascending aorta hypoplasia, asplenia sporadic / dextrocardia, abd. SI, PS, l-TGA, DORV, PAPVR, VSD, 108789- 34 M loss 10p15.2 1M, oligo 3082679 3257396 3131406 2650551 3240195 3272797 174717 203 7 PFKP, PITRM1 ASD, absence of renal to 622246 hepatic IVC with azygous cont., Lt. SVC to coronary sinus, Rt. aortic arch unknown / hydrocephalus, 202625- 35 F loss 13q12.11 1M, oligo 19701063 19919228 19701674 19695681 19904299 19961630 218165 142 30 GJB6, CRYL1 meningomyelocoele, 265949 complex CHD sporadic / dextrocardia, abd. SI, arch abnls., l- NPY, MPP6, DFNA5, TGA, DORV, VSD, 7p15.3- 1863326- OSBPL3, CYCS, ?Kartagener syndrome, Lt. 36 M gain 1M, oligo 24017561 26072707 24157536 24033678 26020862 26052791 2055146 973 54 p15.2 2019113 C7orf31, NPVF, ear cochlear abnls., MIR148A maxillary sinusitis, visual, motor & perceptual delays, mild obesity sporadic / dextrocardia, l- TGA, CAVC, VSD, ASD, DORV, abnl. SVC, sub- aortic stenosis, PDA, n/a, n/a, n/a, n/a, n/a, n/a, 37 F loss 12q24.33 1M 129958950 130676992 718000 535 ADGRD1, LINC01257 common atria, hypoplastic validated validated validated validated validated validated LV, absent Rt. SVC, Lt. SVC not communicating with Rt., asplenia, 1+ H J bodies sporadic / ?scimitar syndrome, mesocardia with dextroposition, abd. SI, gut malrotation, hypoplastic Rt. pulmonary 370k, n/a, n/a, n/a, n/a, n/a, n/a, TTC21B, SCN1A, artery, Lt. SVC to 38 F gain 2q24.3 166468603 167053304 584701 71 oligo validated validated validated validated validated validated SCN9A, SCN7A coronary sinus, ASD, PAPVR, diaphragmatic hernia, possible bifid Rt. kidney, replaced hepatic artery from superior mesenteric artery n/a, n/a, n/a, n/a, n/a, n/a, sporadic / dextrocardia, 39 M gain 5p12-p11 1M, oligo 45563021 46440100 877079 163 HCN1 validated validated validated validated validated validated abd. SI, PA, DORV, abnl.

SVC, VSD, Rt. aortic arch, duodenal stenosis familial, HLHS, CAVC, PA, AVSD, CoA, interrupted IVC, Rt. sided aortic arch, Rt. kidney ectopia with horizontal 370k, n/a, n/a, n/a, n/a, n/a, n/a, BIRC6, TTC27, 40 M gain 2p22.3 32487194 33138319 651125 83 possible mid. abdomen, oligo validated validated validated validated validated validated LTBP1 midline liver with azygous cont., polysplenia, mild ventricular and subarachnoid space enlargement. sporadic / abd. SI, gut MPZ, SDHC, malrotation, PS, d-TGA, CFAP126, FCGR2A, DORV, PAPVR, IVC on HSPA6, FCGR3A, Lt., Lt. SVC to partially 370k, 200137- 41 M gain 1q23.3 159549008 160021458 159546105 159543968 159746242 159762550 472450 48 36 FCGR2C, FCGR3B, unroofed coronary sinus, oligo 218582 FCGR2B, FCRLA, ASD, mitral atresia, infra- FCRLB, DUSP12, diaphragmatic pulmonary ATF6 vein connection below diaphragm COL27A1, MIR455, 370k, 468283- ORM1, ORM2, 41 gain 9q32 116012776 116495996 116011342 115985132 116479625 116549104 483220 116 30 oligo 563972 AKNA, DFNB31, ATP6V1G1, C9orf91 sporadic / situs ambiguus, dextrocardia, l-TGA, 42 F loss 12q15 oligo not called not called 68025619 67981186 68030515 68031979 not called 4896-50793 not called 12 LYZ VSD, DORV, PS, polysplenia HNRNPCL2, PRAMEF3, PRAMEF22, PRAMEF5, PRAMEF6, PRAMEF8, PRAMEF9, PRAMEF14, PRAMEF18, sporadic / eisenmenger PRAMEF16, syndrome, l-TGA, VSD, 167832- 43 M gain 1p36.21 1M, oligo 13081478 13916196 13749684 13712459 13917516 13978325 834718 439 78 PRAMEF20, complete heart block, 265866 PRAMEF21, gout, diverticulosis, PRAMEF8, syncope PRAMEF7, PRAMEF9, PRAMEF14, PRAMEF19, PRAMEF17, PRAMEF20, PRAMEF21, PDPN, PRDM2 unknown / gut malrotation, caudal regression syndrome: double aortic arch, Rt./Lt. arches encircle tracheoesophageal pedicle and fuse NBR1, TMEM106A, posteriorly, Rt. descending LINC00910, ARL4D, 604358- aorta, Lt. subclavian arises 44 M gain 17q21.31 1M, oligo 38804206 39200215 38586778 38558674 39191136 39191278 396009 141 38 MIR2117, DHX8, 632604 from pulmonary artery (Lt. ETV4, MEOX1, Arch), Rt. and Lt. common SOST, DUSP3 carotid arteries arise from aortic trunk (Rt. Arch), absent ductus arteriosis, unilobated Lt. lung. dilated rectum, small anus, recto- urethral fistula, low-set

ears, flexed wrists, flexed hip joints with pterygia, little muscle mass legs, bilat. talipes, spine: segmentation error C5-T2, vertebral column ends at L4. ribs: Rt. = 11; Lt. = 9. Mother with hypothyroidism (synthroid). (Deceased) sporadic / abd. SI, CAVC, DORV, TAPVR, Rt. aortic arch, hemiazygous cont. of KCNH8, EFHB, intrahepatic IVC., complex not not not 1076047- not 45 M gain 3p24.3 oligo 19526648 19510317 20602695 20622036 51 RAB5A, PP2D1, cyanotic CHD, midline analyzed analyzed analyzed 1111719 analyzed KAT2B, SGOL1 liver with numerous hepatic veins, no visible portal structure, ?asplenia (Deceased) not not not 184414- not PIP5K1B, PRKACG, 45 gain 9q21.11 oligo 70813142 70796115 70997556 70998029 305 analyzed analyzed analyzed 201914 analyzed FXN, TJP2 sporadic / situs ambiguus, abd. SI, HLHS, CoA, AS, BAV, MGA, single ventricle, DORV, AOV, arch abnls., both great arteries arise from RV, PDA, mitral atresia, atypical cor triatriatum, 145990- ZFP42, TRIML2, 46 M loss 4q35.2 1M, oligo 189124414 189299208 189145810 189062658 189291800 189314997 174794 92 5 subvalvular AS, very 252339 TRIML1 hypoplastic LV, RV hypertrophy, dil. bile ducts, sludging in gallbladder, nephrocalcinosis, duplex Lt. Kidney, with mod. hydroureteronephrosis of upper pole

Abbreviations: abd. = abdominal; abnl/abnls. = abnormal/abnormalities; ADHD = attention deficit hyperactivity disorder; AS = aortic stenosis; ASD = atrial septal defect; AV = atrioventricular AVC = atrioventricular canal; AVVR = atrioventricular valve regurgitation; BAV = bicuspid aortic valve; bilat. = bilateral; CAVC = complete atrioventricular canal; CHD = congenital heart defect; cont. = continuation; CNV = copy number variant; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DD = developmental delay; dil. = dilated; DILV = double inlet right ventricle; DORV = double outlet right ventricle; HJ bodies = Howell-Jolly bodies; HLHS = hypoplastic left heart syndrome; IAA = interrupted aortic arch; IVC = inferior vena cava; LA = left atrium; l- TGA = levo-transposition of the great arteries; LPA = left pulmonary artery; Lt. = left; LV = left ventricle; MGA = malposition of great arteries; MPA = main pulmonary artery; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PDA = patent ductus arteriosus; PIT = prolonged isolated thrombocytopenia; PS = pulmonic stenosis, Rt. = right; RA = right atrium; RAA = right aortic arch; RV = right ventricle; SI = situs inversus; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; VSD = ventricular septal defect

Supp. Table 2.10.3. Genes identified in rare coding region CNVs detected in the 225 patient heterotaxy cohort.

Gene Protein Gene ID Locus Associated OMIM Phenotypes ATP-binding cassette, sub-family C ABCC1 4363 16p13.11 (CFTR/MRP), member 1 Arterial calcification, generalized, of infancy 2 (AR) (OMIM:614473) ATP-binding cassette, sub-family C ABCC6 368 16p13.11 pseudoxanthoma elasticum (AR) (OMIM:264800) (CFTR/MRP), member 6 pseudoxanthoma elasticum, forme fruste (AD) (OMIM:177850) ACOXL acyl-CoA oxidase-like 55289 2q13

ADGRD1 adhesion G protein-coupled receptor 283383 12q24.33 (GPR133) D1 AGMO alkylglycerol monooxygenase 392636 7p21.2 (TMEM195) AKNA AT-hook transcription factor 80709 9q32

ANAPC1 anaphase promoting complex subunit 1 64682 2q13

ANKRD34A ankyrin repeat domain 34A 284615 1q21.1

ANKRD35 ankyrin repeat domain 35 148741 1q21.1

ARL4D ADP-ribosylation factor-like 4D 379 17q21.31

ATF6 activating transcription factor 6 22926 1q23.3 Achromatopsia 7 (OMIM:616517) ATPase, H+ transporting, lysosomal ATP6V1G1 9550 9q32 13kDa, V1 subunit G1 BCL2L11 BCL2-like 11 (apoptosis facilitator) 10018 2q13

BIRC6 baculoviral IAP repeat containing 6 57448 2p22.3 BUB1 mitotic checkpoint BUB1 699 2q13 Colorectal cancer with chromosomal instability, somatic serine/threonine kinase C16orf45 chromosome 16 open reading frame 45 89927 16p13.11

C9orf91 chromosome 9 open reading frame 91 203197 9q32

CCDC138 coiled-coil domain containing 138 165055 2q13

CD160 CD160 molecule 11126 1q21.1

?Bardet-Biedl syndrome 14 (AR) (OMIM:615991) Joubert syndrome 5 (AR) (OMIM:610188) CEP290 centrosomal protein 290kDa 80184 12q21.32 Leber congenital amaurosis 10 (OMIM:611755) Meckel syndrome 4 (AR) (OMIM:611134) Senior-Loken syndrome 6 (AR) (OMIM:610189) CFAP126 cilia and flagella associated protein 257177 1q23.3 (C1orf192) 126 cholinergic receptor, nicotinic, alpha 7 CHRNA7 1139 15q13.3 Schizophrenia, neurophysiologic defect in (OMIM:118511) (neuronal)

COL27A1 collagen, type XXVII, alpha 1 85301 9q32 ?Steel syndrome (AR) (OMIM:615155) CPPED1 calcineurin-like phosphoesterase 55313 16p13.12 (FLJ11151) domain containing 1 CRYL1 crystallin, lambda 1 51084 13q12.11

CYCS cytochrome c, somatic 54205 7p15.3 Thrombocytopenia 4 (AD) (OMIM:612004) DFNA5 deafness, autosomal dominant 5 1687 7p15.2 Deafness, autosomal dominant 5 (AD) (OMIM:600994) DFNB31 deafness, autosomal recessive 31 25861 9q32 Deafness, autosomal recessive 31 (AR) (OMIM:607084) DEAH (Asp-Glu-Ala-His) box DHX8 1659 17q21.31 polypeptide 8 DOCK8 dedicator of cytokinesis 8 81704 9p24.3 Hyper-IgE recurrent infection syndrome, autosomal recessive (AR) (OMIM:243700) DUSP12 dual specificity phosphatase 12 11266 1q23.3

DUSP3 dual specificity phosphatase 3 1845 17q21.31

Ectodermal dysplasia 10A, hypohidrotic/hair/nail type, autosomal dominant (AD) (OMIM:129490) EDAR ectodysplasin A receptor 10913 2q12.3 Ectodermal dysplasia 10B, hypohidrotic/hair/tooth type, autosomal recessive (AR) (OMIM:224900) Hair morphology 1, hair thickness (OMIM:612630) EFHB EF-hand domain family, member B 151651 3p24.3

ETV4 ets variant 4 2118 17q21.31

FBLN7 fibulin 7 129804 2q13

Lupus nephritis, susceptibility to (AD) (OMIM:152700) Fc fragment of IgG, low affinity IIa, FCGR2A 2212 1q23.3 Malaria, severe, susceptibility to (OMIM:611162) receptor (CD32) Pseudomonas aeruginosa, susceptibility to chronic infection by, in cystic fibrosis (AR) (OMIM:219700) Fc fragment of IgG, low affinity IIb, Malaria, resistance to (OMIM:611162) FCGR2B 2213 1q23.3 receptor (CD32) Systemic lupus erythematosus, susceptibility to (AD) (OMIM:152700) Fc fragment of IgG, low affinity IIc, FCGR2C 9103 1q23.3 Thrombocytopenic purpura, autoimmune (AD) (OMIM:188030) receptor for (CD32) Fc fragment of IgG, low affinity IIIa, FCGR3A 2214 1q23.3 Immunodeficiency 20 (AR) (OMIM:615707) receptor (CD16a) Fc fragment of IgG, low affinity IIIb, FCGR3B 2215 1q23.3 Neutropenia, alloimmune neonatal receptor (CD16b) FCRLA Fc receptor-like A 84824 1q23.3

FCRLB Fc receptor-like B 127943 1q23.3

FOPNL FGFR1OP N-terminal like 123811 16p13.11 (C16orf63) FSTL5 follistatin-like 5 56884 4q32.2

FXN frataxin 2395 9q21.11 Friedreich ataxia / Friedreich ataxia with retained reflexes (AR) (229300) GRIP and coiled-coil domain GCC2 9648 2q12.3 containing 2 Deafness, autosomal dominant 3B (AD) (OMIM:612643) GJB6 gap junction protein, beta 6, 30kDa 10804 13q12.11 Deafness, autosomal recessive 1B (AR) (OMIM:612645) Deafness, digenic GJB2/GJB6 (DD, AR) (OMIM:220290)

Ectodermal dysplasia 2, Clouston type (AD) (OMIM:129500)

gonadotropin-releasing hormone (type GNRHR2 114814 1q12-q21.3 2) receptor 2 GPALPP1 GPALPP motifs containing 1 55425 13q14.12 (KIAA1704) (general transcription factor IIF, GTF2F2 2963 13q14.12-q14.13 polypeptide 2, 30kDa hyperpolarization activated cyclic HCN1 348980 5p12 Epileptic encephalopathy, early infantile, 24 (AD) (OMIM:615871) nucleotide gated potassium channel 1 heterogeneous nuclear HNRNPCL2 440563 1p36.21 ribonucleoprotein C-like 2 HSPA6 heat shock 70kDa protein 6 (HSP70B') 3310 1q23.3

IRC6 baculoviral IAP repeat containing 6 57448 2p22.3

ITGA10 integrin, alpha 10 8515 1q21.1

KN motif and ankyrin repeat domains KANK1 23189 9p24.3 Cerebral palsy, spastic quadriplegic, 2 (OMIM:612900) 1 KAT2B K(lysine) acetyltransferase 2B 8850 3p24.3

potassium channel, voltage gated eag KCNH8 131096 3p24.3 related subfamily H, member 8 potassium channel tetramerization KCTD4 386618 13q14.12 domain containing 4 KH domain containing, RNA binding, KHDRBS2 202559 6q11.1 signal transduction associated 2 KIAA0430 KIAA0430 9665 16p13.11

LIM and senescent cell antigen-like LIMS1 3987 2q12.3 domains 1 LIM and senescent cell antigen-like LIMS3 96626 2q13 domains 3 long intergenic non-protein coding 10013058 LINC00910 17q21.31 RNA 910 1 long intergenic non-protein coding LINC01257 116437 12q24.33 RNA 1257 LIX1L limb and CNS expressed 1 like 128077 1q21.1

LOC440895 two pore channel 3 pseudogene 440895 2q13

latent transforming growth factor beta LTBP1 4052 2p22.3 binding protein 1 LYZ lysozyme 4069 12q15 Amyloidosis, renal (AD) (OMIM:105200) MEOX1 mesenchyme homeobox 1 4222 17q21.31 Klippel-Feil syndrome 2 (AR) (OMIM:214300) MEOX2 mesenchyme homeobox 2 4223 7p21.2

MERTK MER proto-oncogene, tyrosine kinase 10461 2q13 Retinitis pigmentosa 38 (AR) (OMIM:613862) MIR148A microRNA 148a 406940 7p15.2

10031377 MIR2117 microRNA 2117 17q21.31 9 MIR4435-2HG MIR4435-2 host gene 541471 2q13 MIR455 microRNA 455 619556 9q32

MIR484 microRNA 484 619553 16p13.11

membrane protein, palmitoylated 6 MPP6 51678 7p15.3 (MAGUK p55 subfamily member 6) Charcot-Marie-Tooth disease, dominant intermediate D (AD) (OMIM:607791) Charcot-Marie-Tooth disease, type 1B (AD) (OMIM:118200) Charcot-Marie-Tooth disease, type 2I (AD) (OMIM:607677) MPZ myelin protein zero 4359 1q23.3 Charcot-Marie-Tooth disease, type 2J (AD) (OMIM:607736) Dejerine-Sottas disease (AD, AR) (OMIM:145900) Neuropathy, congenital hypomyelinating (AD, AR) (OMIM:605253) Roussy-Levy syndrome (AD) (OMIM:180800) myosin, heavy chain 11, smooth MYH11 4629 16p13.11 Aortic aneurysm, familial thoracic 4 (AD) (OMIM:132900) muscle NBR1 neighbor of BRCA1 gene 1 4077 17q21.31

?Microhydranencephaly (AR) (OMIM:605013) NDE1 nudE neurodevelopment protein 1 54820 16p13.11 Lissencephaly 4 (with microcephaly) (AR) (OMIM:614019) NOMO3 NODAL modulator 3 408050 16p13.11

NPTX2 neuronal pentraxin II 4885 7q22.1

NPVF neuropeptide VF precursor 64111 7p15.3

NPY neuropeptide Y 4852 7p15.3

nudix (nucleoside diphosphate linked NUDT17 200035 1q21.1 moiety X)-type motif 17 nuclear fragile X mental retardation NUFIP1 26747 13q14.12 protein interacting protein 1 ORM1 orosomucoid 1 5004 9q32

ORM2 orosomucoid 2 5005 9q32

OSBPL3 oxysterol binding protein-like 3 26031 7p15.3

Parkinson disease, juvenile, type 2 (AR) (OMIM:211980) Adenocarcinoma of lung, somatic (OMIM:167000) PARK2 parkin RBR E3 ubiquitin protein ligase 5071 6q26 Adenocarcinoma, ovarian, somatic (OMIM:600116) Leprosy, susceptibility to (OMIM:607572) PDPN podoplanin 10630 1p36.21

PDZK1 PDZ domain containing 1 5174 1q21.1

PEX11B peroxisomal biogenesis factor 11 beta 8799 1q21.1 Peroxisome biogenesis disorder 14B (OMIM:614920) PFKP phosphofructokinase, platelet 5214 10p15.2

PIAS3 protein inhibitor of activated STAT, 3 10401 1q21.1

phosphatidylinositol-4-phosphate 5- PIP5K1B 8395 9q21.11 kinase, type I, beta PITRM1 pitrilysin metallopeptidase 1 10531 10p15.2

polymerase (RNA) III (DNA directed) POLR3C 10623 1q21.1 polypeptide C (62kD) polymerase (RNA) III (DNA directed) POLR3GL 84265 1q21.1 polypeptide G (32kD)-like PP2D1 chromosome 7 open reading frame 31 136895 7p15.3 (C7orf31) PP2D1 protein phosphatase 2C-like domain 151649 3p24.3 (C3orf48) containing 1 PRAMEF14 PRAME family member 14 729528 1p36.21

PRAMEF16 PRAME family member 16 654348 1p36.21

PRAMEF17 PRAME family member 17 391004 1p36.21

PRAMEF19 PRAME family member 19 645414 1p36.21 (PRAMEF18) PRAMEF20 PRAME family member 20 645425 1p36.21

PRAMEF21 PRAME family member 21 391001 1p36.21

PRAMEF22 PRAME family member 22 653606 1p36.21

PRAMEF3 PRAME family member 3 401940 1p36.21

PRAMEF5 PRAME family member 5 343068 1p36.21

PRAMEF6 PRAME family member 6 440561 1p36.21

PRAMEF7 PRAME family member 7 441871 1p36.21

PRAMEF8 PRAME family member 8 391002 1p36.21

PRAMEF9 PRAME family member 9 343070 1p36.21

PR domain containing 2, with ZNF PRDM2 7799 1p36.21 domain protein kinase, cAMP-dependent, PRKACG 5568 9q21.11 ?Bleeding disorder, platelet-type, 19 (AR) (OMIM:616176) catalytic, gamma RAB5A, member RAS oncogene RAB5A 5868 3p24.3 family RANBP2 RAN binding protein 2 5903 2q12.3 Encephalopathy, acute, infection-induced, 3, susceptibility to (AD) (OMIM:608033) RBM8A RNA binding motif protein 8A 9939 1q21.1 Thrombocytopenia-absent radius syndrome (AR) (OMIM:274000) RANBP2-like and GRIP domain RGPD4 285190 2q12.3 containing 4 RGPD4 antisense RNA 1 (head to RGPD4-AS1 729121 2q12.3 head) RNF115 ring finger protein 115 27246 1q21.1

SCN1A sodium channel, voltage gated, type I 6323 2q24.3 Dravet syndrome (AD) (OMIM:607208)

alpha subunit Epilepsy, generalized, with febrile seizures plus, type 2 (AD) / Febrile seizures, familial, 3A (AD) (OMIM:604403) Migraine, familial hemiplegic, 3 (AD) (OMIM:609634) sodium channel, voltage gated, type SCN7A 6332 2q24.3 VII alpha subunit Epilepsy, generalized, with febrile seizures plus, type 7 (AD) / Febrile seizures, familial, 3B (AD) (OMIM:613863) Erythermalgia, primary (AD) / Small fiber neuropathy (AD) (OMIM:133020) sodium channel, voltage gated, type IX SCN9A 6335 2q24.3 HSAN2D, autosomal recessive (AR) / Insensitivity to pain, congenital (AR) (OMIM:243000) alpha subunit Paroxysmal extreme pain disorder, (AD) (OMIM:167400) Dravet syndrome, modifier of (AD) (OMIM:607208) succinate dehydrogenase complex, Gastrointestinal stromal tumor (IC, AD) (OMIM:606764) SDHC subunit C, integral membrane protein, 6391 1q23.3 Paraganglioma and gastric stromal sarcoma (OMIM:606864) 15kDa Paragangliomas 3 (AD) (OMIM:605373) sema domain, immunoglobulin domain SEMA3A (Ig), short basic domain, secreted, 10371 7q21.11 Hypogonadotropic hypogonadism 16 with or without anosmia (OMIM:614897) (semaphorin) 3A SEPT10 septin 10 151011 2q13

SGOL1 shugoshin-like 1 (S. pombe) 151648 3p24.3 Chronic atrial and intestinal dysrhythmia (AR) (OMIM:616201) SH3RF3 SH3 domain containing ring finger 3 344558 2q13

10028721 SH3RF3-AS1 SH3RF3 antisense RNA 1 2q12.3 6 solute carrier family 5 (sodium/choline SLC5A7 60482 2q12.3 Neuronopathy, distal hereditary motor, type VIIA (AD) (OMIM:158580) cotransporter), member 7 SNX29 sorting nexin 29 92017 16p13.13

Craniodiaphyseal dysplasia, autosomal dominant (AD) (OMIM:122860) SOST sclerostin 50964 17q21.31 Sclerosteosis 1 (AR) (OMIM:269500) Van Buchem disease (AR) (OMIM:239100) SOWAHC sosondowah ankyrin repeat domain 65124 2q13 (ANKRD57) family member C ST6 beta-galactosamide alpha-2,6- ST6GAL2 84620 2q12.2-q12.3 sialyltranferase 2 sulfotransferase family, cytosolic, 1C, SULT1C2 6819 2q12.3 member 2 sulfotransferase family, cytosolic, 1C, SULT1C3 442038 2q12.3 member 3 sulfotransferase family, cytosolic, 1C, SULT1C4 27233 2q12.3 member 4 Cholestasis, progressive familial intrahepatic 4 (AR) (OMIM:615878) TJP2 tight junction protein 2 9414 9q21.11 Hypercholanemia, familial (OMIM:607748) TMEM106A transmembrane protein 106A 113277 17q21.31

TMEM132B transmembrane protein 132B 114795 12q24.31

TMEM87B transmembrane protein 87B 84910 2q13

TRIML1 tripartite motif family-like 1 339976 4q35.2

TRIML2 tripartite motif family-like 2 205860 4q35.2

transfer RNA glutamic acid (anticodon 10012651 TRNAE-UUC 1p36.13 UUC) 7 TTBK2 tau tubulin kinase 2 146057 15q15.2 Spinocerebellar ataxia 11 (AD) (OMIM:604432) Nephronophthisis 12 (OMIM:613820) TTC21B tetratricopeptide repeat domain 21B 79809 2p24.3 Short-rib thoracic dysplasia 4 with or without polydactyly (AR) (OMIM:613819) TTC27 tetratricopeptide repeat domain 27 55622 2p22.3

TXNIP thioredoxin interacting protein 10628 1q21.1

ubiquitin protein ligase E3 component UBR1 197131 15q15.2 Johanson-Blizzard syndrome (AR) (OMIM:243800) n-recognin 1 VPS13B vacuolar protein sorting 13 homolog B 157680 8q22.2 Cohen syndrome (AR) (OMIM:216550) Desbuquois dysplasia 2 (AR) (OMIM:615777) XYLT1 xylosyltransferase I 64131 16p12.3 Pseudoxanthoma elasticum, modifier of severity of (AR) (OMIM:264800) ZC3H6 zinc finger CCCH-type containing 6 376940 2q14.1

ZC3H8 zinc finger CCCH-type containing 8 84524 2q14.1

ZFP42 ZFP42 zinc finger protein 132625 4q35.2

ZNF480 zinc finger protein 480 147657 19q13.41

ZNF528 zinc finger protein 528 84436 19q13.41

ZNF534 zinc finger protein 534 147658 19q13.41

ZNF578 zinc finger protein 578 147660 19q13.41

ZNF610 zinc finger protein 610 162963 19q13.41

ZNF804A zinc finger protein 804A 91752 2q32.1

ZNF880 zinc finger protein 880 400713 19q13.41

Abbreviations: AD = autosomal dominant; AR = autosomal recessive; DD = digenic dominant; IC = isolated cases; OMIM = Online Mendelian Inheritance in Man (http://www.omim.org/)

Supp. Table 2.10.4. Rare copy number variants at loci previously associated with CHD.

Size Size Agilent Genes at locus CHD associations at Locus Patient Gender Type Platform Illumina (min-max) Affected Genes Inheritance / Phenotype associated References locus1,2 (bp) (bp) with CHD

TXNIP, POLR3GL, ANKRD34A, LIX1L, RBM8A, 1q21.1 deletion Klopocki, et al., 2007; not 300679- GNRHR2, PEX11B, ITGA10, sporadic / levocardia, situs (Thrombocytopenia- 1q21.1 19 M gain oligo RBM8A Brunetti-Pierri, et al., analyzed 960808 ANKRD35, PIAS3, NUDT17, ambiguus, abd. SI, PS, l-TGA, absent radius syndrome, 2008 POLR3C, RNF115, CD160, RBM8A) (274000) PDZK1 ST6GAL2, RGPD4-AS1, RGPD4, SLC5A7, SULT1C3, SULT1C2, SULT1C4, GCC2, not 3683400- sporadic / dextrocardia, ASD, PA, Wenger, et al., 2004; 2q12.2-q13 gain oligo LIMS1, RANBP2, CCDC138, analyzed 3832914 atrial inversion, common AV Rudd, et al., 2009; EDAR, SH3RF3-AS1, 2q13-q23 duplication, valve, single RV; asplenia, history TMEM87B, Cooper, et al., 2011; 29 M SH3RF3, SEPT10, SOWAHC, 2q13 duplication of multiple sinus infections, FBLN7 Soemedi, et al., 2012; LIMS3, LOC440895 2q13 deletion ADHD Yu, et al., 2012; BUB1, ACOXL, BCL2L11, Russell, et al., 2014 not 1695755- MIR4435-2HG, ANAPC1, 2q13 gain oligo analyzed 2585889 MERTK, TMEM87B, FBLN7, ZC3H8, ZC3H6 sporadic / scimitar syndrome, mesocardia with dextroposition, abd. SI, gut malrotation, hypoplastic Rt. PA, Lt. SVC to 2q24.3-2q32.1 Nimmakayalu, et al., TTC21B, SCN1A, SCN9A, 2q24.3 38 F gain 370k, oligo 584701 n/a, validated coronary sinus, ASD, PAPVR, duplication, TTC21B 2012; Lim, et al., SCN7A diaphragmatic hernia, possible 2q24 deletion 2014 bifid Rt. kidney, replaced hepatic artery from superior mesenteric artery familial (1st cousin with renal McLeod, et al., 1990; anomaly) / SI totalis, TAPVR, arch Hopkin, et al., 1997; abnls, Rt. aortic arch., dextrocardia not 208827- Stevenson, et al., 6q26 25 M loss oligo PARK2 with atrioventricular and 6q26 deletion analyzed 426776 2004; Eash, et al., ventricularterial concordance, 2005; Dobyns, et al., hemivertebrae, microtia 2008 (Deceased). familial / situs ambiguus, ASD, AVVR, single ventricle, gut 7p21.1 deletion Megarbane, et al., not 67216- malrotation, liver abnls., dysplastic (Saethre-Chotzen TWIST1, 7p21.1 27 M loss oligo MEOX2, AGMO 2001; Fryssira, et al., analyzed 114770 single AV valve, TGA with aorta Syndrome, TWIST1) MEOX2 2011 anterior and pulmonary artery (101400) posterior, enlarged RA familial (male sib expired with same defects) / situs ambiguus, 8p22.2 deletion, arch abnls., d-TGA, DORV, VSD, Balikova, et al., 2009; duplication (Cohen 261043- ASD, anterior & rightward aorta, Rivera-Brugues, et al., 8q22.2 26 M loss 1M, oligo 286743 VPS13B (COH1) Syndrome, COH1) COH1 271615 severe valvar/subvalvar PS, small 2011; Concolino, et (216550) MPA, atretic LPA, RAA, Rt. ear al., 2012 8q22.2-24.3 duplication lobe "pushed up", foot positional defects sporadic / situs ambiguus, abd. SI, 15q13.3 duplication, Sharp, et al., 2008; 15q13.3 18 F gain 1M, oligo 579670 n/a, validated CHRNA7 gut malrotation, single ventricle, 15q13.3 deletion van Bon, et al., 2009;

PS, MGA, DILV, TAPVR, (612001) Masurel-Paulet, et al., PAPVR, bilat. SVC, VSD, PDA, 2010; Szafranski, et asplenia, midline liver, cleft al., 2010; Tropeano, et lip/palate al., 2014 C16orf45, KIAA0430, NDE1, Kuang, et al., 2011; 16p13.11 duplication, 16p13.11- not 2282653- MIR484, MYH11, FOPNL, sporadic / levocardia, situs MYH11, Nagamani, et al., 19 M gain oligo 16p13.11 deletion p12.3 analyzed 2900749 ABCC1, ABCC6, NOMO3, ambiguus, abd. SI, PS, l-TGA, NOMO3 2011; Warburton, et (132900) XYLT1 al., 2014 unknown / gut malrotation, caudal regression syndrome: double aortic arch, Rt./Lt. arches encircle tracheoesophageal pedicle and fuse posteriorly, Rt. descending aorta, Lt. subclavian arises from PA (Lt. arch), Rt. and Lt. common carotid 17q21.31-q21.32 Koolen, et al., 2006; arteries arise from aortic trunk (Rt. NBR1, TMEM106A, duplication, Koolen, et al., 2008; arch), absent ductus arteriosis. 604358- LINC00910, ARL4D, 17q21.31 deletion Tan, et al., 2009; 17q21.31 44 M gain 1M, oligo 396009 unilobated Lt. lung. dilated rectum, ARL4D 632604 MIR2117, DHX8, ETV4, (Koolen-de Vries Kitsiou-Tzeli, et al., small anus, recto-urethral fistula, MEOX1, SOST, DUSP3 syndrome, KANSL1 2012; Priest, et al., low-set ears, flexed wrists, flexed (610443) 2012 hip joints with pterygia, little muscle mass legs, bilat. talipes, spine: segmentation error C5-T2, vertebral column ends at L4. ribs: Rt.= 11; Lt.= 9. Mother with hypothyroidism (synthroid). (Deceased)

1OMIM reference numbers and causative genes are indicated whenever possible 2 Literature review was completed for both duplications and deletions at each locus. Only CHD-associated abnormalities are listed. Searches were carried out on Pubmed and Google Scholar using the format “locus + qualifiers”, where “locus” represents the locus of interest and “qualifiers” encompasses one or more of the following search terms: deletion, duplication, congenital heart defects, CHDs, heart, cardiac, cardiovascular. Follow-up searches were carried out to determine reported gene-specific associations with CHD. Listed references are not intended to be comprehensive, but provide a representative sample of available literature. Abbreviations: abd. = abdominal; abnl/abnls. = abnormal/abnormalities; ADHD = attention deficit hyperactivity disorder; ASD = atrial septal defect; AV = atrioventricular; AVVR = atrioventricular valve regurgitation; bilat. = bilateral; CHD = congenital heart defect; CNV = copy number variant; d-TGA = dextro-transposition of the great arteries; DILV = double inlet right ventricle; DORV = double outlet right ventricle; l-TGA = levo-transposition of the great arteries; LPA = left pulmonary artery; Lt. = left; MPA = main pulmonary artery; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PS = pulmonic stenosis, Rt. = right; RA = right atrium; RAA = right aortic arch; RV = right ventricle; SI = situs inversus; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; TGA = transposition of the great arteries; VSD = ventricular septal defect

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Supp. Table 2.10.5. Rare CNVs identified at loci associated with known microdeletion / duplication syndromes but not heterotaxy or CHD.

Includes Size Size Smallest reported affected Agilent Affected Locus Patient Gender Type Platform Illumin Inheritance / Phenotype microduplication / gene(s) Key Features Reference (min-max) Genes a (bp) deletion at locus from Htx (bp) cohort? sporadic / situs ambiguus, abd. SI, gut malrotation, single ventricle, DF, thin/sparse hair, FD, 2517224- Rosenfeld, et 2q32.1 17 M gain 370k 2778870 ZNF804A PS, MGA, DILV, TAPVR, 2q32-33.1 deletion No CL/P, LD, GR, genital 2580386 al., 2009 PAPVR, bilat. SVC, VSD, PDA, abnormalities asplenia, midline liver, CL/P variable phenotypes reported: sporadic / HLHS, CoA, arch abnls., Not 7q22.1-q22.3 dysmorphic features, brain Al-Hassnan, et 7q22.1 31 F loss oligo 98000 NPTX2 d-TGA, DORV, ASD, mitral atresia No called deletion abnls., finger abnls., genital al., 2011 with severely hypoplastic LV abnls., DD, MR Yes 202625- GJB6, unknown / hydrocephalus, Lagou, et al., 13q12.11 35 F loss 1M, oligo 218165 13q12.11 deletion (GJB6, DF, DD 265949 CRYL1 meningomyelocoele, complex CHD 2014 CRYL1) Duplication: minor DF, short stature, abnl. ears, short neck, Yes MR, DD, seizures ZNF480, (ZNF480, ZNF610, 19q13.33-ter Mikelsaar, et sporadic / dextrocardia, PA, l-TGA, ZNF610, Deletion: DF, severe MR, 120782- ZNF880, deletion al., 2001; 19q13.33 42 F loss 1M, oligo 162000 VSD, ASD, arrhythmia ZNF880, DD, absence of speech, 162530 ZNF528, 19q13.33 Carvalheira, et (Pacemaker), scoliosis ZNF528, severe muscle spasticity, ZNF534, duplication al., 2014 ZNF534, scoliosis, short stature, ZNF578 ZNF578) microcephaly, teeth malocclusion, undeveloped secondary sex characteristics

Abbreviations: abd. = abdominal; abnl. = abnormal; ASD = atrial septal defect; bilat. = bilateral; CAVC = complete atrioventricular canal; CHD = congenital heart defect; CL/P = cleft lip and palate; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DD = developmental delay; DILV = double inlet right ventricle; DORV = double outlet right ventricle; FD = fetal demise; GR = growth retardation; HLHS = hypoplastic left heart syndrome; Htx; heterotaxy; IVC = inferior vena cava; LD = learning difficulties; l-TGA = levo-transposition of the great arteries; LV = left ventricle; MGA = malposition of great arteries; MR = mental retardation; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PDA = patent ductus arteriosus; PS = pulmonary stenosis; Rt. = right; SI = situs inversus; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; VSD = ventricular septal defect

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CHAPTER 3: Genetic and functional analyses of ZIC3 variants in congenital heart disease

Adapted and used with permission from “Cowan et al., Genetic and functional analyses of ZIC3 variants in congenital heart disease, Human Mutation, 35(1): 66-75, DOI: 10.1002/humu.22457,

Available online: http://onlinelibrary.wiley.com/doi/10.1002/humu.22457/pdf.

3.1. ABSTRACT

Mutations in zinc-finger in cerebellum 3 (ZIC3) result in heterotaxy, a condition characterized by abnormal organ laterality, complex cardiovascular malformations, and isolated

CHD. The majority of reported mutations cluster in zinc-finger domains. We previously demonstrated that many of these lead to aberrant ZIC3 subcellular trafficking. A relative paucity of N- and C-terminal mutations has, however, prevented similar analyses in these regions.

Notably, an N-terminal polyalanine expansion was recently identified in a patient with

VACTERL, suggesting a potentially distinct function for this domain. Here, we report ZIC3 sequencing results from 440 unrelated patients with heterotaxy and CHD, the largest cohort yet examined. Variants were identified in 3.8% of sporadic cases, exceeding previous estimates of

1%. Eight of 11 were novel, including 5 N-terminal variants. Subsequent functional analyses included 4 additional reported but untested variants. Aberrant cytoplasmic localization and decreased luciferase transactivation were observed for all zinc-finger variants, but not for downstream or in-frame upstream variants, including both analyzed polyalanine expansions.

Collectively, these results expand the ZIC3 mutational spectrum, support a higher than expected prevalence in sporadic cases, and suggest alternative functions for terminal mutations, highlighting a need for further study of these domains.

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3.2. INTRODUCTION

Bilateral symmetry is broken early in development following establishment of the anterior-posterior and dorsal-ventral body axes. Embryos of some species begin to manifest left- right (L-R) asymmetries as early as initial cleavage stages, establishing molecular gradients and initiating signaling cascades that foreshadow additional L-R pattering processes at the ciliated embryonic node (reviewed inVandenberg and Levin, 2010). Failure to properly establish L-R asymmetry results in heterotaxy, a condition of significant morbidity and mortality associated with abnormal positioning and disrupted morphogenesis of the thoraco-abdominal viscera. Much of the clinical severity results from complex and heterogeneous heart malformations that are classically accompanied by abnormalities in spleen position and number, malrotation of the developing gut tube, and/or situs anomalies of the liver and stomach. As discussed in the previous chapters, the overall genetics of heterotaxy are complex and have been reviewed in detail elsewhere (Sutherland and Ware, 2009); however, both de novo and familial mutations have been reported, including pedigrees suggestive of autosomal recessive, autosomal dominant, and X-linked inheritance patterns (Vitale, et al., 2001; Belmont, et al., 2004; Zhu, et al., 2006).

The clinical significance of the zinc-finger in cerebellum 3 (ZIC3) gene (OMIM

#300265), the first human heterotaxy gene to be described (Gebbia, et al., 1997) and the only known genetic cause of X-linked heterotaxy (OMIM #306955), is well documented(Gebbia, et al., 1997; Megarbane, et al., 2000; Ware, et al., 2004; Ware, et al., 2006a; Chhin, et al., 2007; De

Luca, et al., 2010; Tzschach, et al., 2010; Wessels, et al., 2010; Chung, et al., 2011;

D'Alessandro, et al., 2011; Cast, et al., 2012). Estimates from our initial analyses of 194 individuals with heterotaxy and heterotaxy spectrum CHD suggested that 75% of pedigrees consistent with X-linked inheritance and 1% of sporadic heterotaxy cases are attributable to ZIC3

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mutations (Ware, et al., 2004). Nevertheless, variable expressivity is common, resulting in a wide spectrum of clinical manifestations ranging from situs abnormalities and isolated heart defects to more complex diagnoses involving additional midline, gastrointestinal, urogenital, and/or central nervous system anomalies (Casey, et al., 1993; Gebbia, et al., 1997; Ware, et al.,

2004; De Luca, et al., 2010; Wessels, et al., 2010; Chung, et al., 2011; D'Alessandro, et al., 2011;

Cast, et al., 2012). Such phenotypic complexity is mirrored in mice deficient for Zic3, which exhibit a wide variety of congenital abnormalities including heterotaxy, vertebral and rib anomalies, neural tube defects, and cerebellar hypoplasia (Purandare, et al., 2002; Ware, et al.,

2006b; Jiang, et al., 2013; Sutherland, et al., 2013).

Recent reports additionally associate ZIC3 mutations with VACTERL, a constellation of defects (vertebral, anal, cardiac, tracheo-esophageal, renal, radial, and limb) demonstrating phenotypic overlap with heterotaxy (Wessels, et al., 2010; Chung, et al., 2011). A polyalanine tract expansion in one of these patients (Wessels, et al., 2010) is particularly intriguing as similar tract expansions have been associated with a variety of human genetic disorders, including holoprosencephaly caused by mutations in ZIC2 (Messaed and Rouleau, 2009).

Both the complexity of ZIC3-associated phenotypes and its broad expression during embryogenesis (Nagai, et al., 1997; McMahon and Merzdorf, 2010; Quinn, et al., 2012) are indicative of diversity in ZIC3 developmental function, including recognized roles in neural and neural crest development (Nakata, et al., 1997; Klootwijk, et al., 2000), limb bud digitation

(Quinn, et al., 2012), cardiac morphogenesis, and L-R patterning (Kitaguchi, et al., 2000;

Purandare, et al., 2002; Ware, et al., 2004; Ware, et al., 2006a; Ware, et al., 2006b; Zhu, et al.,

2007b; Zhu, et al., 2007a; Jiang, et al., 2013; Sutherland, et al., 2013). Studies utilizing Zic3-null mice indicate that Zic3 is required for progression into and through gastrulation (Ware, et al.,

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2006b; Cast, et al., 2012) and that it acts upstream of Nodal signaling at the embryonic node

(Jiang, et al., 2013; Sutherland, et al., 2013).

The ZIC3 gene encodes a highly conserved zinc-finger protein belonging to the GLI superfamily of transcription factors (Mizugishi, et al., 2001; Sakai-Kato, et al., 2008). The most highly conserved elements of the GLI superfamily members are the zinc finger DNA-binding domains (Herman and El-Hodiri, 2002; Aruga, et al., 2006). Five of these domains encompass over a third of the ZIC3 amino acid sequence and collectively comprise a DNA-binding domain that facilitates not only transcriptional activation of target genes, but also binding of transcriptional cofactors and subcellular trafficking of the ZIC3 protein (Mizugishi, et al., 2001;

Ware, et al., 2004; Bedard, et al., 2007; Hatayama, et al., 2008; Zhu, et al., 2008; Lim, et al.,

2010). Interestingly, relatively few ZIC3 polymorphisms have been reported in 1000 genomes

(1KG, http://browser.1000genomes.org/) and NHLBI Exome Sequencing Project (ESP, http://evs.gs.washington.edu/EVS/) databases, potentially reflecting high levels of sequence conservation throughout the entire coding sequence (Ware, et al., 2004).

In this study, we report results from ZIC3 mutation screening in 440 unrelated patients with heterotaxy and isolated heterotaxy-spectrum CHD and perform in vitro functional testing of

15 ZIC3 variants. These studies represent not only the first major sequencing effort to include coverage of the recently identified fourth exon of ZIC3 (Bedard, et al., 2011), but also the first functional assessment of expansions of the ZIC3 polyalanine tract. Our sequencing results increase the total number of reported ZIC3 variants from 23 to 31 (Stenson, et al., 2003) and notably include the most N-terminal (Asp6GlufsX32) and C-terminal (Ala447Gly) variants yet identified. Collectively, these data expand the known ZIC3 mutation spectrum, particularly in N- terminal encoding sequences, and reveal a higher than anticipated incidence in patients with

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sporadic heterotaxy (3.8% vs. 1%), particularly in affected males. Furthermore, our functional analyses confirm the importance of the zinc-finger regions for both ZIC3 localization and transcriptional target activation and suggest the existence of alternative functional consequences for variants in N- and C-terminal regions, including those in the polyalanine tract.

3.3. MATERIALS AND METHODS

3.3.1. Patient recruitment and phenotypic classification

Informed consent was obtained from all study participants according to protocols approved by the institutional review board at Cincinnati Children's Hospital Medical Center

(CCHMC). Using our previously described classification system (Ware, et al., 2004), each participant was categorized as having SI, heterotaxy, or isolated heterotaxy spectrum CHD based on phenotypic information collected from patient histories and chart review. Heterotaxy was considered to be “familial” if the presenting pedigree met one or more of the following criteria:

1) autosomal dominant, autosomal recessive, or X-linked inheritance; 2) more than one family member with heterotaxy or laterality disorder regardless of degree of relationship; 3) heterotaxy in the proband and a first degree relative with isolated CHD; and/or 4) heterotaxy in the proband and a first degree relative with situs-related defect(s), including but not limited to gut malrotation, isolated dextrocardia, bilateral superior vena cava, or right-sided stomach.

Heterotaxy identified in pedigrees not meeting one or more of these criteria was classified as

“sporadic”.

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3.3.2. Variant detection

Genomic DNA was prepared from participant blood samples following standard protocols. DNA was PCR-amplified with oligonucleotide primers designed to encompass all

ZIC3 coding regions and splice junctions (RefSeq: NM_003413.3), including the recently identified fourth exon (Bedard, et al., 2011). Sequencing primer sequences were as follows: exon

1aF: 5’-TCCTATCCCTCTGCAGGAGAC-3’ and exon 1aR 5’-

ACTCGCGCGTTGAGTTGAAGG-3’; exon 1bF 5’-GAATCCCTTCGGGGACTCAAC-3’ and exon 1bR 5’-CAGCTCCTGCTTGATAGGCTG-3’; exon 1cF 5’-

CAGTTTCCTAACTACAGCCCC-3’ and exon 1cR 5’-GTGGATGGTCACTGACAGCGC-3’; exon 2F 5’-GCTGCTTGCCTCTGAGAAAC-3’ and exon 2R 5’-

ACGTGGAAGACAGTGGGTTG-3’, exon 3F 5’-GCTCTTGTTTTTGCTTGCAC-3’ and exon

3R 5’-CATTTCCATCTGATTGGTCTC-3’; and exon 4F 5’-

AAGAGGAAATGTGGCCTGTTT-3’ and exon 4R 5’-CACTTCAAGGTAACAGACATCCA-

3’. All amplifications were performed at 57°C as 20L reactions containing 20ng genomic DNA,

10L Fast Master Mix (Roche) and 10pmol of each primer. PCR-amplified products were treated with exonuclease 1 and shrimp alkaline phosphatase prior to sequencing. Bi-directional sequencing was performed using BigDye Terminator v3.1 on an ABI 3730XL DNA analyzer and mutation analysis was completed with Mutation Surveyor v4.0.5 (SoftGenetics Inc.). Detected variants were then queried against the Human Gene Mutation Database (HGMD, http://www.hgmd.cf.ac.uk/) and nomenclature was assigned to novel changes according to

Human Genome Variation Society (HGVS, http://www.hgvs.org/) guidelines.

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3.3.3. Variant interpretation

Pathogenicity of each ZIC3 variant was interpreted according to 2014 clinical molecular diagnostic laboratory standards taking into account type of mutation, bioinformatic prediction of protein function (if applicable), prevalence in control populations (1KG, ESP public variant databases), segregation with disease phenotype, and functional analyses.

3.3.4. Expression constructs and mutagenesis:

Methods for generating an N-terminal hemagglutinin (HA)-tagged, wild-type ZIC3 expression construct (HA-ZIC3) have been previously described (Ware, et al., 2004). This plasmid contains the entire human ZIC3-A ORF subcloned into a pHM6 expression vector

(Roche). Mutations were introduced into the wild-type ZIC3 sequence via Quik-Change site- directed mutagenesis (Agilent) and confirmed by Sanger sequencing. Primers used to generate each mutant ZIC3 construct are listed in Supp. Table 3.9.1.

3.3.5. Cell culture

NIH/3T3 cells were obtained from the American Type Culture Collection. Cells were grown in high glucose DMEM (HyClone) supplemented with 10% newborn calf serum

(HyClone) using a Forma Series II 3110 Water-Jacketed CO2 Incubator (Thermo Scientific) maintained at 37°C and 5% CO2.

3.3.6. Luciferase assays

Luciferase assays were conducted as previously described (Ware, et al., 2004). NIH/3T3 cells were seeded onto 9.6cm2 6-well plates (BD-Falcon) at 2 X 105 cells/ml one day prior to transfection with FuGENE HD transfection reagent (Promega) following manufacturer’s

123 protocols. A total of 2g of DNA and a DNA:FuGENE HD ratio of 1:3 were used for each transfection. Wild-type or mutant HA-ZIC3 constructs were co-transfected with an SV40 firefly luciferase reporter (pGL3-SV40, Promega), and a Renilla luciferase reporter (pRL-TK, Promega) to control for transfection efficiency. A promoterless luciferase vector (pGL3-Basic, Promega) was used for normalization. Cells were lysed 24 h after transfection and luciferase activities were determined using the Dual Luciferase Reporter Assay System (Promega) and a Glomax 96

Microplate Luminometer (Promega). Firefly luciferase activities were normalized to Renilla luciferase activity. Fold activations for each ZIC3 mutant were calculated with respect to wild- type values. Results for each construct represent average values obtained from a minimum of 3 independent experiments repeated in triplicate. Statistical analyses were completed using

GraphPad Instat v. 3.

3.3.7. Immunocytochemistry and subcellular localization

NIH/3T3 cells were seeded onto 15mm diameter circular glass coverslips (Electron

Microscopy Sciences) at 1 X 105 cells/ml one day prior to transfection with either wild-type or mutant HA-ZIC3. For each transfection, 2g of DNA were transfected at a 1:3 ratio with

FuGENE HD transfection reagent (Promega). After 24 h, transfected cells were washed in PBS and fixed for 20 minutes at room temperature in 4% paraformaldehyde/PBS. Following 3 washes in PBS and 3 additional washes in 0.1% BSA/PBS, cells were blocked for 1 h at room temperature with 50% normal donkey serum/0.3% Triton X-100/PBS. Cells were subsequently incubated for 1 h at room temperature in a 1:200 dilution of anti-HA rabbit polyclonal antibody

(NB600-363, Novus Biologicals), washed 3 times in 0.1% BSA/PBS, incubated for 1 h in a

1:200 dilution of Alexa Fluor 488 goat anti-rabbit IgG (A-11008, Molecular Probes), washed 3

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more times in 0.1% BSA/PBS, and mounted using Vectashield Hardset with DAPI (Vector

Labs). Slides were imaged using an Eclipse E400 fluorescence microscope equipped with a

DXM1200F digital camera and ACT-1 v. 2.62 imaging software (Nikon). Merged images were generated in Adobe Photoshop CS5 extended v. 12.0x32 and subsequently imported to Image J v. 1.45s (rsbweb.nih.gov/ij/) for analysis. Cells were scored as having either “nuclear”,

“cytoplasmic”, or “nuclear and cytoplasmic” ZIC3 expression. At least 250 cells across a minimum of 3 independent experiments were scored for each ZIC3 construct.

3.4. RESULTS

3.4.1. Identification of ZIC3 variants

ZIC3 coding and splice junction regions were fully sequenced in 440 unrelated patients with heterotaxy or isolated heterotaxy-spectrum CHD. The demographics of the cohort are summarized in Table 3.7.1. A total of 11 ZIC3 variants were identified in 15 patients, including 8 that were novel. Phenotypic characteristics of each patient are summarized in Table 3.7.2.

Collectively, the identified variants encompass 7 point mutations (2 nonsense, 5 missense), 2 indels (1 deletion, 1 deletion-insertion), and 2 duplications (including 1 polyalanine expansion).

Figure 3.8.1. (panel a) depicts amino acid and cDNA locations for each variant. Truncations and their predicted effects on ZIC3 protein structure are also shown (Figure 3.8.1, panel b). In all but one pedigree, ZIC3 sequencing was performed on the proband. The sole exception was an asymptomatic mother (Patient 12) whose DNA was sequenced in the absence of an available sample from her son with heterotaxy. Sequencing identified a heterozygous C to A transversion resulting in a nonsense mutation that prematurely truncates the ZIC3 protein at a highly conserved serine 252 residue bordering the first zinc-finger domain (Ser252X). The severe

125 phenotype of the proband and the carrier status of his mother imply likely X-linked transmission of the variant allele. Pathogenicity of the variant is also supported by the identification of a nearby nonsense mutation (Gln249X) in multiple individuals of a family with comparably complex CHD and situs abnormalities (Ware, et al., 2004).

Familial disease was associated with 4 of the 11 ZIC3 variants. Two of these encode truncated proteins. The first, an out-of-frame indel at the 5’ end of the ZIC3 gene

(Asp6GlufsX32), was associated with heterotaxy in a male proband (Patient 1) and his maternal half-brother. The resulting protein is severely truncated at 36 amino acids and lacks all known functional domains. The second variant, an out-of-frame duplication of a single G nucleotide near the 3’ end of exon 1, was identified in a male proband with heterotaxy and a family history of 2 siblings with Ivemark syndrome, one of which also had CHDs (Patient 13). This variant was predicted to encode a 342 amino acid protein missing all but the first zinc-finger domain

(Glu291GlyfsX53). Notably, a nonsense mutation resulting in loss of the same functional regions

(Gln294X) was previously reported in the context of familial heterotaxy (Gebbia, et al., 1997).

The 2 remaining familial variants identified in the cohort were missense. The first, an N-terminal variant replacing a semi-conserved proline with an alanine residue (Pro217Ala), was identified in a female proband with heterotaxy (Patient 11) and was inherited from her asymptomatic mother.

The family history was significant for a stillborn brother of unknown cause and a mutation- positive sister with CHD. The second identified missense mutation replaces a highly conserved histidine residue with an asparagine in the second zinc-finger domain (His318Asn) and was found in a female proband with isolated CHD (Patient 14). The family history was exceptional for a brother with situs abnormalities and a maternal uncle with complex CHD. A nearby and

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similarly conserved zinc-finger 2 missense mutation (Thr323Met) was previously identified in over 10 related individuals with heterotaxy and related anomalies (Gebbia, et al., 1997).

A total of 8 ZIC3 variants were identified in individuals with sporadic disease, including the aforementioned Ser252X nonsense mutation in Patient 12. These variants ranged from single amino acid substitutions to more complex genetic abnormalities. Five were point mutations, including 4 missense variants found in 3 males and in a single female with isolated CHD. Each was found either upstream or downstream of the zinc-finger domains. Two (Gly17Cys and

Ala447Gly) affected highly conserved amino acids, while 2 (Ser109Cys and Pro217Ala) altered amino acids incompletely conserved among species. Although Ser109Cys and Ala447Gly are novel, both Gly17Cys and Pro217Ala were previously reported in patients with isolated CHD

(Ware, et al., 2004; De Luca, et al., 2010). In the cohort, Gly17Cys and Pro217Ala variants were also identified in 3 patients with laterality defects (Patients 3, 5, and 11), including a patient with

Williams syndrome (Patient 4). The remaining sporadic point mutation, a novel nonsense mutation encoding a 154 amino acid protein truncated upstream of the zinc-finger regions

(Glu155X), was identified in a male patient with heterotaxy (Patient 8). Additionally, an out-of- frame deletion removing 16bp midway through exon 1 (Tyr199delfsX19), was detected in a male patient with heterotaxy (Patient 9). The resulting 216 amino acid protein lacks all 5 zinc-finger domains. Finally, a male patient with isolated CHD (Patient 6) was found to carry a single alanine residue expansion (Ala53dup) in the ZIC3 polyalanine tract. A similar single alanine expansion was recently identified in a male patient with isolated CHD (d-TGA) , while larger expansions of 2 and 3 alanine residues were previously identified in a male patient with

VACTERL association (Wessels, et al., 2010) and a female patient with heterotaxy, respectively

(D'Alessandro, et al., 2013).

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In summary, we have identified 11 ZIC3 variants in a cohort of 440 patients with heterotaxy and CHD, including 8 that were novel, 5 of which fall in the N-terminal domain.

Detection of variants in 10/264 (3.8%) sporadic cases, including 8/154 (5.2%) males and 2/111

(1.8%) females, indicated a higher than expected yield patients with sporadic disease, based on past estimates of 1% (Ware, et al., 2004). Our analyses additionally verify previous associations of ZIC3 mutations with isolated CHD (Megarbane, et al., 2000; Ware, et al., 2004; D'Alessandro, et al., 2013) and reinforce existing evidence for pathogenicity of ZIC3 deficiency in a subset of carrier females (Gebbia, et al., 1997; Ware, et al., 2004; De Luca, et al., 2010; Chung, et al.,

2011; Cast, et al., 2012).

3.4.2. ZIC3 variants alter luciferase reporter gene transactivation

As a transcription factor, ZIC3 function is dependent on both its localization to the nucleus and its subsequent ability to activate target genes. Previous studies have demonstrated the importance for the zinc-finger domains in fulfilling these functions (Ware, et al., 2004;

Bedard, et al., 2007; Hatayama, et al., 2008); however, the impact of mutations in upstream and downstream regions, including the polyalanine tract, remain less clearly defined. We therefore sought to assess the pathogenic significance of each of the 11 ZIC3 variants identified in the cohort. To maximize the breadth of our analyses, we supplemented these with 4 recently published variants for which functional assessments were lacking. Site-directed mutagenesis was used to generate mutant ZIC3 constructs from a previously described expression vector encoding the wild-type ZIC3-A open reading frame fused to an N-terminal hemagglutinin (HA) epitope tag

(Ware, et al., 2004). Wild-type and mutant ZIC3 constructs were subsequently co-transfected into NIH/3T3 cells with an SV40 promoter-driven firefly luciferase reporter (Ware, et al., 2004).

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In agreement with previous studies (Ware, et al., 2004; Bedard, et al., 2007; Hatayama, et al.,

2008), ZIC3 mutations in the zinc-finger regions significantly disrupted reporter gene transactivation relative to wild-type controls (Figure 3.8.2). Included in this group were 2 missense variants affecting highly conserved amino acid residues (His318Asn, Arg350Gly), as well as 6 truncating variants individually producing proteins missing at least 4 out of 5 zinc- fingers (Asp6GlufsX32, Ala50ProfsX9, Glu155X, Tyr199delfsX19, Ser252X,

Glu291GlyfsX53). Of the remaining variants, Ala447Gly was exceptional, demonstrating significant and reproducible increase in luciferase reporter transactivation. As the most C- terminal ZIC3 variant identified to date, Ala447Gly lies downstream of any mapped functional domains and affects only the larger ZIC3-A isoform, which preferentially splices in the

Ala447Gly-containing exon 3 at the expense of the ZIC3-B specific exon 4 (Bedard, et al., 2011).

3.4.3. Zinc-finger domain mutations demonstrate aberrant cytoplasmic subcellular localization

As an additional assessment of the functional impact of ZIC3 variants, transient transfections of HA-tagged ZIC3 constructs and immunofluorescence analyses were performed to determine the effect of each variant on ZIC3 protein subcellular localization. Localization experiments for the current study were carried out in the same NIH/3T3 cells used for the luciferase transactivation studies. Cells were counted and scored as having “nuclear”,

“cytoplasmic” or “nuclear and cytoplasmic” ZIC3, as described (see Materials and Methods).

Results from these experiments are summarized in Figure 3.8.3.

GLI superfamily protein subcellular localization has previously been shown to be cell line dependent (Koyabu, et al., 2001). In NIH/3T3 cells, exogenously expressed ZIC3 localizes to the nucleus in approximately 50-55% of cells with the remaining 45-50% demonstrating

129 mixed nuclear and cytoplasmic ZIC3 localization (Figure 3.8.3). Similar localization patterns were observed following transfection with the Gly17Cys construct, encoding the most N- terminal missense mutation reported to date (this study, De Luca, et al., 2010). Slightly fewer cells (40-45%) exhibited nuclear localization with either of the polyalanine expansions or with the C-terminal Ser402Pro variant; each of which lie downstream from Gly17Cys in non-zinc- finger domains. Three similarly positioned missense mutations (Ser109Cys, Pro217Ala,

Ala447Gly) also yielded large proportions of cells with nuclear ZIC3 (30-35%), albeit at decreased levels relative to wild-type transfections. In contrast, ZIC3 was exclusively nuclear in

<20% of cells transfected with mutant constructs affecting the zinc-finger domains (Figure

3.8.3.O). Notably, each of these 5 variants (Glu155X, Ser252X, Glu291GlyfsX53, His318Asn,

Arg350Gly) are predicted to disrupt one or more amino acids in previously mapped nuclear localization (NLS) and nuclear export (NES) signals (Hatayama, et al., 2008; Bedard, et al.,

2011). Most striking was His318Asn, which affects a single highly conserved histidine residue in overlapping zinc finger 2 NLS and NES sequences (Bedard, et al., 2007; Hatayama, et al., 2008).

Finally, it should be noted that no detectable fluorescent signal was observed for 2 out-of-frame mutations (Asp6GlufsX32 and Ala50ProfsX9), presumably due to the extremely small sizes of each predicted protein product (Figure 3.8.1.B). Results from all functional assays are summarized in Table 3.7.3. Collectively, they support previous findings that ZIC3 subcellular localization and transcriptional activity is most sensitive to mutations affecting amino acids in zinc-finger domains and suggest the likelihood of alternative pathogenic mechanisms for variants affecting the N- and C-terminal domains (Ware, et al., 2004; Bedard, et al., 2007; Hatayama, et al., 2008).

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3.5. DISCUSSION

To date, sequencing efforts have identified 23 novel mutations affecting ZIC3 coding regions (Stenson, et al., 2003; D'Alessandro, et al., 2013), establishing ZIC3 loss of function as an important cause of both familial (~75%) and sporadic (~1%) heterotaxy (Ware, et al., 2004).

In the present study, we report sequencing results from an additional 440 patients with heterotaxy and/or heterotaxy-spectrum CHD, reassess mutation prevalence, and describe functional analyses for 15 previously uncharacterized ZIC3 variants, 8 of which were novel to the cohort.

In agreement with previous findings (Gebbia, et al., 1997; Ware, et al., 2004; De Luca, et al., 2010; Chung, et al., 2011; Cast, et al., 2012), ZIC3 variants were detected in both males and females with familial and sporadic disease. Notably, the observed 3.8% (10/264) incidence in sporadic cases was higher than expected based on our previous analyses of a similar patient population (Ware, et al., 2004). The majority of this increase is attributable to a higher than anticipated proportion of mutation carriers among males with sporadic disease (8/153 = 5.2%).

Pathogenicity determinations and relevant data for all ZIC3 variants are summarized in Table

3.7.3 and reflect clinical molecular diagnostic laboratory standards for mutation interpretation

(see Materials and Methods). Ten of the 15 analyzed ZIC3 variants were interpreted as disease causing mutations, including 4 missense mutations (His318Asn, Arg350Gly, Ser402Pro,

Ala447Gly), 2 nonsense mutations (Glu155X, Ser252X) and 4 out-of-frame mutations predicted to cause premature protein truncation upstream of all zinc finger domains (Asp6GlufsX32,

Ala50ProfsX9, Tyr199delfsX19, Glu291GlyfsX53 ). Each mutation was absent from 1KG and

ESP public databases, was predicted to be pathogenic by relevant predictive software algorithms, and disrupted ZIC3 subcellular localization and nuclear activity in vitro. Two additional

131 missense variants (Ser109Cys, Pro217Ala) were identified in <1% of individuals in 1KG and

ESP control populations and yielded milder, but still abnormal in vitro functional results alongside less damaging in silico pathogenicity predictions. An increased prevalence of the

Pro217Ala variant has been observed among ostensibly healthy individuals of African American descent [MAFAA = 0.02 (EVS), 0.01 (1KG)]; however, our functional results suggest that

Pro217Ala and Ser109Cys have the potential to act as susceptibility alleles alongside other genetic or environmental modifying factors. Lastly, one missense mutation (Gly17Cys) and both polyalanine expansions (Ala53dup, Ala52_Ala53dup) were interpreted to be of uncertain pathogenic significance based on lack of effect on tested functional parameters. Future experimentation will be required to definitively determine the pathogenic potential of these variants; however, support for potential pathogenicity of Gly17Cys is provided by the fact that

Gly17Cys affects a highly conserved residue, was predicted to be functionally deleterious by 4/5 mutation pathogenicity programs, and is represented in <1% of 1KG and ESP control populations.

Although pathogenic mutations have been identified throughout the ZIC3 gene, most have clustered in the latter portion of exon 1, a sequence encoding 3 of 5 highly conserved

Cys2His2 zinc-finger domains. Pathogenic significance of this DNA-binding region has been demonstrated by cell-based analyses of human heterotaxy mutations (Ware, et al., 2004;

Hatayama, et al., 2008) and by observations of L-R patterning defects in Xenopus embryos expressing zinc-fingerless ZIC3 protein (Kitaguchi, et al., 2000). Furthermore, recent studies have identified NLS and NES signals overlapping amino acids in these regions (Bedard, et al.,

2007; Hatayama, et al., 2008), indicating their importance for not only DNA-binding, but also

ZIC3 subcellular trafficking.

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In contrast to ‘classical’ NLSs, which comprise one or more short stretches of basic amino acids (reviewed in Lange, et al., 2007), the ZIC3 NLS is formed through juxtaposition of physically separated (interspersed) basic residues throughout the zinc finger domain (Hatayama, et al., 2008). Similar interspersed NLSs have been described in other zinc-finger transcription factors (Pandya and Townes, 2002). Systematic mutagenesis of ZIC3 coding sequences has mapped the functional ZIC3 NLS to a long stretch of amino acids (292-356) in zinc-fingers 2 + 3

(ZF2+3) (Hatayama, et al., 2008) and to a bipartite sequence (367-382,403-412) in zinc-fingers 4 and 5 (ZF4+5) (Bedard, et al., 2007). Surprisingly, the ZF2+3 NLS was also found to overlap a cryptic CRM-1 dependent NES spanning amino acids 313-325 (Bedard, et al., 2007). Structural changes in ZIC3 protein structure, including those caused by heterotaxy mutations, can expose the export motif and facilitate cytoplasmic localization of properly imported ZIC3 protein

(Bedard, et al., 2007).

We have previously reported significant decreases in ZIC3 nuclear localization and luciferase reporter transactivation for both missense (His286Arg, Thr323Met) and nonsense

(Cys268X, Gln292X) mutations affecting the ZF2+3 NLS (Ware, et al., 2004). In the present study, a missense mutation in zinc-finger 2 (His318Asn), as well as several truncation mutants lacking these and all other downstream zinc-finger domains (Glu155X, Ser252X,

Glu291GlyfsX53), resulted in similarly severe functional abnormalities. The His318Asn mutation uniquely maps to amino acids overlapping both the ZF2+3 NLS and the cryptic NES, which may account for the striking lack of nuclear ZIC3 in nearly 75% of His318Asn-transfected cells (Figure 3.8.3). In contrast, ZIC3 nuclear localization and luciferase transactivation levels were considerably less affected by non-truncating mutations in regions outside of NLS/NES regions, including those upstream of (Gly17Cys, Ala53dup, Ala52_Ala53dup, Ser109Cys,

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Pro217Ala), between (Ser402Pro), and downstream of (Ala447Gly) the ZF2+3 and ZF4+5 localization signals (Ware, et al., 2004, present study). Collectively, these analyses underline the functional relevance of ZIC3 mutations affecting the zinc-finger, NLS, and NES domains and lend support to heterotaxy pathogenesis models whereby altered subcellular trafficking of mutant

ZIC3 prevents downstream transcriptional activation of as yet unidentified developmentally critical genes (Ware, et al., 2004; Bedard, et al., 2007; Hatayama, et al., 2008).

The N- and C-terminal domains of the ZIC3 protein are highly conserved, but have yet to be ascribed specific cellular functions. Only 9 N-terminal variants have been previously reported, including an out-of-frame deletion (Ala50ProfsX9) (Cast, et al., 2012), 3 polyalanine tract expansions (Wessels, et al., 2010; D'Alessandro, et al., 2013), a pair of nonsense mutations

(Ser43X, Gln249X) (Ware, et al., 2004), and 3 missense mutations (Ala33Val, Gly17Cys, and

Pro217Ala) (Ware, et al., 2004; De Luca, et al., 2010; D'Alessandro, et al., 2013). Three of these variants (Gly17Cys, Ala53dup, Pro217Ala) were additionally identified in patients from our cohort. We have significantly expanded this mutation spectrum through identification of 5 additional N-terminal variants (Asp6GlufsX32, Ser109Cys, Glu155X, Tyr199delfsX19, and

Ser252X). Collectively, 7 of the 14 known N-terminal variants (Gly17Cys, Ala33Val,

Ser109Cys, Pro217Ala, and the 3 polyalanine expansions) preserve the ZIC3 reading frame and, presumably, maintain structural and functional integrity of all downstream zinc-finger sites.

Functional analyses indicate that amino acids in the N-terminal domain, including those in the polyalanine tract, have little to no effect on ZIC3 subcellular trafficking and transcriptional activation. Although it remains a possibility that the identified N-terminal variants lack functional significance, high levels of amino acid conservation throughout the domain suggest the potential for alternative pathogenic mechanism(s). It is noteworthy that the polyalanine

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expansions, if definitely established as pathogenic, would represent the shortest disease-causing mutations of this type yet reported.

A comparison of phenotypic characteristics among ZIC3 mutation carriers and non- carriers in our heterotaxy and CHD cohort revealed few significant differences (Supp. Table

3.9.2), with the exception of an over-representation of pulmonary atresia in the ZIC3 mutation cohort (40% vs. 15.29%, p = 0.0215, F.E.). In general, in-frame N-terminal ZIC3 variants were associated with isolated and sporadic CHD (Patients 2, 4, 6, 7, and 10). These findings echo previously published data for Gly17Cys and Pro217Ala variants (Ware, et al., 2004; De Luca, et al., 2010) and suggest potentially milder phenotypes for N-terminal mutations relative to those in zinc-finger regions, which are more often associated with familial heterotaxy, situs inversus, and/or other situs abnormalities (Gebbia, et al., 1997; Ware, et al., 2004; Chhin, et al., 2007;

Wessels, et al., 2010; D'Alessandro, et al., 2011; Cast, et al., 2012, this study). These findings are also in agreement with existing in vitro data, which almost invariably indicates more severe abrogation of ZIC3 function by mutations in zinc-finger domains (Ware, et al., 2004; Hatayama, et al., 2008; D'Alessandro, et al., 2013; this study). Nevertheless, more severe phenotypes in 3 patients with N-terminal mutations (Patients 3, 5, and 11) and recent identification of polyalanine tract expansions in patients with VACTERL and heterotaxy (Wessels, et al., 2010; D'Alessandro, et al., 2013) suggest that definitive genotype-phenotype correlations are unlikely to be so simplistic and may be influenced by other modifying factors. Future genetic and functional studies will undoubtedly help to further delineate the phenotypic and functional consequences of

N-terminal variants of uncertain significance, including those affecting the polyalanine tract.

We recently reported the existence of a second, shorter ZIC3 splice variant, which preferentially includes a downstream fourth exon at the expense of exon 3 (Bedard, et al., 2011).

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Consequently, the C-terminal domain is alternatively encoded by exon 3 (in ZIC3-A) or 4 (in

ZIC3-B), depending on post-transcriptional splicing. Our mutation analyses represent the first broad scale sequencing efforts to include full sequencing coverage of exon 4, although no ZIC3-

B specific mutations were identified in our cohort. We did, however, identify a novel C to G transversion in exon 3, resulting in substitution of a highly conserved alanine for a glycine

(Ala447Gly) in a female patient with isolated CHD. This mutation uniquely and reproducibly increased activation of the luciferase reporter, despite a 10% decrease in cells with nuclear localization of ZIC3 (Figure 3.8.2). As of the time of writing, no other C-terminal missense mutations have been reported, precluding genotype-phenotype comparisons. The functional relationship between ZIC3-A and ZIC3-B remains uncertain, with both isoforms expressed in similar tissues and at similar time-points during embryonic development (Bedard, 2011).

Nevertheless, although both ZIC3-A and ZIC3-B are capable of transcriptionally activating a Gli- binding site reporter, only ZIC3-A does so synergistically with Gli3, suggesting some level of functional distinction between the two isoforms (Bedard, 2011). Because all subcellular localization and luciferase reporter analyses were carried out using the ZIC3-A ORF, we cannot exclude the possibility that individual mutations may have different effects on ZIC3-B function.

Future analyses utilizing both isoforms will, consequently, be beneficial in determining the pathophysiological relevance N- and C-terminal variants of uncertain functional impact, as well as the extent to which activating mutations like Ala447Gly contribute to CHD and heterotaxy.

In summary, we have described in vitro functional significance of 15 ZIC3 variants, including 8 novel variants identified in a cohort of 440 patients with heterotaxy and related CHD.

Twelve of these 15 variants were considered to be disease causing or disease associated, while 3 variants, including both analyzed polyalanine tract expansions, remain of uncertain significance.

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Collectively, these analyses greatly expand the known ZIC3 mutation spectrum, particularly in the relatively less characterized N-terminal protein domain, and suggest that ZIC3 mutations are more common in sporadic heterotaxy than previously anticipated. This information is essential for providing accurate risk-based counseling to families with heterotaxy and associated cardiovascular disease. Finally, our data confirm the importance of the zinc-finger domains for

ZIC3 cellular trafficking and nuclear function, and suggest the potential existence of alternative pathogenic mechanisms for variants affecting N- and C-terminal regions, including the polyalanine tract.

3.6. ACKNOWLEDGEMENTS

We thank the patients and families for their participation. This work was supported by

National Institutes of Health grants RO1 HL088639 and HL088639-03S1 (to S. Ware.).

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3.7. TABLES

Table 3.7.1. Heterotaxy and CHD cohort demographics.

Gender Inheritance Racial and Ethnic Categories Totals (%) Females Males Familial Sporadic Unknown American Indian/Alaskan 1 (0.23) 0 1 0 1 0

Asian 14 (3.18) 5 9 1 10 3

Black or African American 52 (11.82) 23 29 7 30 15

Caucasian 222 (50.45) 93 129 32 126 64

Hispanic / Latino 77 (17.50) 26 51 8 57 12

Native Hawaiian / Other Pacific Islander 1 (0.23) 0 1 0 1 0

Mixed 23 (5.23) 9 14 6 17 0

Other1 6 (1.36) 2 4 0 6 0

Unknown 44 (10.00) 23 21 5 16 23

Totals (%) 440 (100.00) 181 (41.14) 259 (58.86) 59 (13.41) 264 (60.00) 117 (26.59)

Percentages are in parentheses and relative to the 440 patient total. 1Includes Arabic, Sephardic Jewish, and Sri-Lankan racial categories

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Table 3.7.2. Summary of ZIC3 variants detected in heterotaxy cohort.

Amino Acid Ethnicity / Patient Nucleotide Change Exon Gender Inheritance Phenotype Detailed Phenotype Change Race Caucasian / Familial; maternal half- 1 p. Asp6GlufsX32 c.18_19delinsA 1 M African brother with asplenia (d. age Heterotaxy Gut malrotation, asplenia, ASD, CAVC, PA American 7 years) HLHS, PS, l-TGA, DILV, ASD, MS with 2 p.Gly17Cys c.49G>T 1 M Caucasian Sporadic Isolated CHD hypoplastic MV, dysplastic PV, congenital coronary anomaly 3 p.Gly17Cys c.49G>T 1 M Caucasian Unknown Heterotaxy Esophageal atresia Sporadic; sister and mother Williams Syndrome, dextrocardia, d-TGA,TA, 4 p.Gly17Cys c.49G>T 1 F Unknown mutation positive, presumed Isolated CHD severely hypoplastic RV, mild hypoplasia of normal phenotype ascending Ao Dextrocardia, Abd SI, single ventricle, PA, 5 p.Gly17Cys c.49G>T 1 M Caucasian Sporadic SI totalis TA, PAPVR, dilated Ao, R Ao arch, MPA w/o connection, R spleen, L liver, ASD, VSD Hispanic or 6 p.Ala53dup c.157_159dupGCC 1 M Sporadic Isolated CHD PS, d-TGA, VSD Latino African 7 p.Ser109Cys c.326C>G 1 M Sporadic Isolated CHD d-TGA, PDA American Dextrocardia, CAVC, midline liver, PA, Ao 8 p.Glu155X c.463G>T 1 M Caucasian Sporadic Heterotaxy arch abnormalities, MGA, DORV, TAPVR, SVC abnormalities, VSD, ASD c. Dextrocardia, VSD, ventriculomegaly, no 9 p. Tyr199delfsX19 delTACCGCCCAGT 1 M Unknown Sporadic Heterotaxy stomach GGCCA595-610 Hispanic or Ventricular inversion, hypoplastic LV & MV, 10 p.Pro217Ala c.649C>G 1 M Sporadic Isolated CHD Latino d-TGA, DORV, VSD Familial; maternally African inherited; mutation-positive Gut ,malrotation, Polysplenia, midline liver, 11 p.Pro217Ala c.649C>G 1 F Heterotaxy American sister with CHDs; stillborn interrupted IVC, HCM brother, cause unknown Unaffected carrier mother; sequenced in 12 p.Ser252X c.755C>A 1 F Caucasian Sporadic lieu of sample from Abd SI, asplenia, CAVC, PA, VSD, ASD affected son with heterotaxy Familial; 2 sibs with 13 p.Glu291GlyfsX53 c.866dupG 1 M Caucasian “Ivemark”, 1 with heart Heterotaxy Abd SI, asplenia, CAVC, PA, d-TGA defect Familial; brother with VSD, L hip dysplasia 14 p.His318Asn c.952C>A 1 F Caucasian isomerism/asplenia, mat. VACTERL-like

uncle with complex CHD 15 p.Ala447Gly c.1340C>G 3 F Caucasian Sporadic Isolated CHD PA, VSD, ASD

1 RefSeq: NP_003404.1, NM_003413.3 Abbreviations: Abd SI = abdominal situs inversus; Ao = aorta; ASD = atrial septal defect; CAVC = complete atrioventricular canal; CHD = congenital heart disease; d-TGA = dextro-transposition of

139 the great arteries; DILV = double inlet left ventricle; DORV = double outlet right ventricle; HCM = hypertrophic cardiomyopathy, HLHS = hypoplastic left heart syndrome; IVC = inferior vena cava; l- TGA = levo-transposition of the great arteries; LV = left ventricle; MGA = malposed great arteries; MPA = main pulmonary artery; MS = mitral stenosis; MV = mitral valve; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PDA = patent ductus arteriosus; PS = pulmonic stenosis; PV = pulmonary valve; RV = right ventricle; SI = situs inversus; SVC = superior vena cava; TA = tricuspid atresia; TAPVR = total anomalous pulmonary venous return; VACTERL = vertebral, anomalies, anal atresia, cardiovascular anomalies, tracheoesophageal fistula, renal/radial anomalies, limb defects; VSD = ventricular septal defect

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Table 3.7.3. Summary of variant analyses.

1KG / ESP Pathogenicity Reporter Gene Subcellular 3 Variant Variant Type 1 2 Variant Interpretation Reference (MAF) Predicitons Transactivation Localization p. Asp6GlufsX32 Frameshift na / na na Decreased na Disease causing This study (c.18_19delinsA) p.Gly17Cys Variant of uncertain This study; Missense 0.002 / 0.003 + + + + - Normal Normal (c.49G>T) significance DeLuca et al. 2010 p. Ala50ProfsX9 Frameshift na / na na Decreased na Disease causing Ma et al. 2011 (c.148delG) p.Ala53dup Polyalanine Variant of uncertain This study; na / na na Normal Normal (c.157_159dupGCC) expansion significance D’Alessandro et al. 2013 p.Ala52_Ala53dup Polyalanine Variant of uncertain na / na na Normal Normal Wessels et al. 2010 (c.154_159dupGCCGCC) expansion significance p.Ser109Cys Missense na / 0.0002 + + na - - Normal Abnormal Likely disease associated This study (c.326C>G) p.Glu155X Nonsense na / na na Decreased Abnormal Disease causing This study (c.463G>T) p.Tyr199delfsX19 (c.595-610del Frameshift na / na na Decreased Abnormal Disease causing This study TACCGCCCAGTGGCCA)

This study; p.Pro217Ala Missense 0.003 / 0.008 - - + - + Normal Abnormal Likely disease associated D’Alessandro et al. 2013; (c.649C>G) Ware et al. 2004

p.Ser252X Nonsense na / na na Decreased Abnormal Disease causing This study (c.755C>A)

p.Glu291GlyfsX53 Frameshift na / na na Decreased Abnormal Disease causing This study (c.866dupG)

p.His318Asn Missense na / na + + + + - Decreased Abnormal Disease causing This study (c.952C>A) p.Arg350Gly Missense na / na + + + + + Decreased Abnormal Disease causing D’Alessandro et al. 2011 (c.1048A>G) p.Ser402Pro Missense na / na + + + + + Normal Abnormal Disease causing Ma et al. 2011 (c.1204T>C) p.Ala447Gly Missense na / na + + + + - Increased Abnormal Disease causing This study (c.1340C>G) 1MAFs represent allele frequencies for the minor allele (in all populations) from the 1000 Genomes Project (1KG, http://www.1000genomes.org/) and NHLBI Exome Sequencing Project (ESP, http://evs.gs.washington.edu/EVS/) databases. 1KG data encompass whole genome indels and SNP variants from 1092 individuals representing 14 world-wide populations (20110521-V3), while ESP data encompass exome SNP variants from 6503 individuals of European American and African American descent (ESP6500SI-V2). 2Missense variant pathogenicity predictions were generated using (from left to right) Polyphen (genetics.bwh.harvard.edu/pph2/), SIFT (sift.jcvi.org/), PANTHER (http://www.pantherdb.org/tools/csnpScoreForm.jsp), Mutation Taster (http://www.mutationtaster.org/), and PMut (http://mmb2.pcb.ub.es:8080/PMut/) algorithms. Variants were predicted to be damaging (+), or benign/neutral (-). Analyses which were not possible are also indicated (na). 3 Based on clinical molecular diagnostic laboratory guidelines (see Materials and Methods)

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3.8. FIGURES

Figure 3.8.1. ZIC3 structure. (A) Wild-type human ZIC3, RefSeq: NP_003404.1, NM_003413.3. Locations of ZIC3 variants are indicated. cDNA numbering begins at the A (position +1) of the ATG initiation codon (codon 1). ZF = zinc-finger domain. (B) Predicted protein structures and amino acid lengths for wild-type and truncating ZIC3 variants. Arrows represent mutation sites. Hatched bars indicate altered out-of-frame amino-acid sequences.

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Figure 3.8.2. Mutations in ZIC3 alter transactivation of an SV40 luciferase reporter. Wild- type (HA-ZIC3 WT) or mutant ZIC3 constructs were co-transfected into NIH/3T3 cells with pGL3-SV40 firefly and pRL-TK renilla luciferase reporters. Luciferase activities were measured 24 hours post-transfection. Mean-fold activations relative to wild-type are shown. Results represent average luciferase activations across a minimum of 4 individual experiments. Standard errors are indicated by vertical lines. Asterisks denote statistical significance (p<0.05) by two- tailed, unpaired Student’s T-tests assuming unequal variance.

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Figure 3.8.3. Mutations in ZIC3 alter subcellular localization. (A-N) Representative subcellular localization for wild-type (WT) or mutant HA-tagged ZIC3 constructs transfected into NIH/3T3 cells. Anti-HA and DAPI staining are shown individually and merged. (O) Cells transfected with each ZIC3 construct were classified as having either nuclear, cytoplasmic, or nuclear and cytoplasmic ZIC3 localization. A minimum of 250 cells were scored for each construct.

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3.9. SUPPLEMENTARY MATERIALS

Supp. Table 3.9.1. Mutagenic primers used to generate HA-tagged variant ZIC3 expression constructs.

Forward Primer Sequence Reverse Primer Sequence Variant (5’  3’) (5’  3’)

p. Asp6GlufsX32 ACCCATGACGATGCTCCTGGAAGAGGCCCGCAGTT GCCCAGGGAACTGCGGGCCTCTTCCAGGAGCATC (c.18_19delinsA) CCCTGGGC GTCATGGGT p.Gly17Cys GGGCTGGGAGTGTGCAGCTTCGGCG CGCCGAAGCTGCACACTCCCAGCCC (c.49G>T) p. Ala50ProfsX9 CACGCCGCCGCCGCCCCGCCGCCGCCGCTGCC GGCAGCGGCGGCGGCGGGGCGGCGGCGGCGTG- (c.148delG) p.Ala53dup ACTCAACCCACGCCGCCGCCGCCGCCGCCGCCGCC GCTCAGCTTGAAGGCAGCGGCGGCGGCGGCGGCG (c.157_159dupGCC) GCCGCTGCCTTCAAGCTGAGC GCGGCGGCGGCGTGGGTTGAGT

p.Ala52_Ala53dup ACTCAACCCACGCCGCCGCCGCCGCCGCCGCCGCC GCTCAGCTTGAAGGCGGCGGCAGCGGCGGCGGCG (c.154_159dupGCCGCC) GCTGCCGCCGCCTTCAAGCTGAGC GCGGCGGCGGCGGCGTGGGTTGAGT p.Ser109Cys ACGGTGGCGCTGCCTGTGCCGCCT AGGCGGCACAGGCAGCGCCACCGT (c.326C>G) p.Glu155X GCTGGCATCCCCTAGCCCCCTAGCT AGCTAGGGGGCTAGGGGATGCCAGC (c.G463T) p.Tyr199delfsX19 (c.595-610del GGCCGTGCTGACCCAGCCCGCGC GCGCGGGCTGGGTCAGCACGGCC TACCGCCCAGTGGCCA) p.Ser252X ATCAAGCAGGAGCTGTAGTGCAAGTGGATCGAC GTCGATCCACTTGCACTACAGCTCCTGCTTGAT (c.755C>A) p.Glu291GlyfsX53 GCATGTGGGGGGGCCCGGAGCAGAA TTCTGCTCCGGGCCCCCCCACATGC (c.866dupG) p.His318Asn AGTACAAACTGGTCAACAACATCCGAGTGCACACG CGTGTGCACTCGGATGTTGTTGACCAGTTTGTACT (c.952C>A) p.Arg350Gly CTCAAGATCCACAAGGGGACCCACACAGGTG CACCTGTGTGGGTCCCCTTGTGGATCTTGAG (c.1048A>G) p.Ser402Pro ACGCACCCGAGCCCCCTGCGCAAAC GTTTGCGCAGGGGGCTCGGGTGCGT (c.1204T>C) p.Ala447Gly CACTAAAACCCCTTCTGGAGTTCAAACTAGCACCA TGGTGCTAGTTTGAACTCCAGAAGGGGTTTTAGTG (c.1340C>G)

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Supp. Table 3.9.2. Phenotypic summary of ZIC3 variant carriers relative to the 440 patient heterotaxy and CHD cohort. Heterotaxy ZIC3 Mutation Non-Mutation Cohort Cohort Cohort % % % Overall Phenotype: n n n p-value1 (n/225) (n/49) (n/176) SI totalis 47 10.68 1 6.67 46 10.82 NS, p = 1.000 Heterotaxy 248 56.36 8 53.33 240 56.47 NS, p = 0.7895 Isolated CHD 117 26.59 5 33.33 112 26.35 NS, p = 0.5570 VACTERL-like 28 6.36 1 6.67 27 6.35 NS, p = 1.000 Abdomen/gastrointestinal:

Abdominal SI 129 29.32 3 20.00 126 29.65 NS, p = 0.5688 Bile duct hypoplasia/biliary atresia 7 1.59 0 0.00 7 1.65 NS, p = 1.000 Gallbladder abnormalities 10 2.27 0 0.00 10 2.35 NS, p = 1.000 Malrotation of the gut 93 21.14 2 13.33 91 21.41 NS, p = 0.7472 Asplenia 112 25.45 3 20.00 109 25.65 NS, p = 0.7696 Polysplenia 46 10.45 1 6.67 45 10.59 NS, p = 1.000 Multilobated spleen 1 0.23 0 0.00 1 0.24 NS, p = 1.000 HJ bodies 1 0.23 0 0.00 1 0.24 NS, p = 1.000 Imperforate anus 14 3.18 0 0.00 14 3.29 NS, p = 1.000 Renal anomalies 24 5.45 0 0.00 24 5.65 NS, p = 1.000 Liver anomalies 24 5.45 1 6.67 23 5.41 NS, p = 0.5749 Skeletal/limbs

Levocardia 20 8.89 4 8.16 16 9.09 1.0000 (NS) Mesocardia 5 2.22 1 2.04 4 2.27 1.0000 (NS) Dextrocardia 13 5.78 2 4.08 11 6.25 0.7388 (NS) Atria:

Common atrium 13 2.95 0 0.00 13 3.06 NS, p = 1.000 Atrial isomerism 9 2.05 0 0.00 9 2.12 NS, p = 1.000 ASD 132 30.00 6 40.00 126 29.65 NS, p = 0.3982 AV canal 98 22.27 3 20.00 95 22.35 NS, p = 1.000 Single ventricle / HLHS 67 15.23 3 20.00 64 15.06 NS, p = 0.7115 VSD 176 40.00 8 53.33 168 39.53 NS, p = 0.2960 Vessels:

SVC abnormality 83 18.86 1 6.67 82 19.29 NS, p = 0.3226 IVC abnormality 93 21.14 1 6.67 92 21.65 NS, p = 0.2113 TAPVR 63 14.32 1 6.67 62 14.59 NS, p = 0.7066 PAPVR 20 4.55 2 13.33 18 4.24 NS, p = 0.1446 Great arteries/conotruncal:

Aortic arch abnormalities 87 19.77 2 13.33 85 20.00 NS, p = 0.7455 d-TGA 116 26.36 5 33.33 111 26.12 NS, p = 0.5546 l-TGA 40 9.09 1 6.67 39 9.18 NS, p = 1.000 DILV 17 3.86 1 6.67 16 3.76 NS, p = 0.4517 DORV 114 25.91 2 13.33 112 26.35 NS, p = 0.3730 PA 71 16.14 6 40.00 65 15.29 S, p = 0.0215 PS 78 17.73 2 13.33 76 17.88 NS, p = 1.000 AS 11 2.50 0 0.00 11 2.59 NS, p = 1.000 CoA 28 6.36 0 0.00 28 6.59 NS, p = 0.6129 BAV 5 1.14 0 0.00 5 1.18 NS, p = 1.000

1 p-values for statistical significance (p<0.05) represent comparisons between ZIC3 mutation and non-mutation cohorts by two-tailed Fischer’s Exact tests using Graphpad Instat v.3 software. NS = non-significant, S = significant. Abbreviations: AS = aortic stenosis; ASD = atrial septal defect; CAVC = complete atrioventricular canal; BAV = bicuspid aortic valve; CHD = congenital heart disease; CoA = coarctation of the aorta; d-TGA = dextro-transposition of the great arteries; DILV = double inlet right ventricle; DORV = double outlet right ventricle; HJ bodies = Howell-Jolly bodies; IVC = inferior vena cava; l-TGA = levo-transposition of the great arteries; PA = pulmonary atresia; PAPVR = partial anomalous pulmonary venous return; PS = pulmonic stenosis, including sub, valvar, and supravalvar but not branch PS; SVC = superior vena cava; TAPVR = total anomalous pulmonary venous return; TOF = Tetralogy of Fallot; VACTERL = vertebral, anal, cardiac, tracheo-esophageal, renal, radial, limb; VSD = ventricular septal defect

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Bedard JE, Purnell JD, Ware SM. 2007. Nuclear import and export signals are essential for proper cellular trafficking and function of ZIC3. Hum Mol Genet 16:187-98.

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D'Alessandro LC, Casey B, Siu VM. 2011. Situs Inversus Totalis and a Novel ZIC3 Mutation in a Family with X-linked Heterotaxy. Congenit Heart Dis

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Koyabu Y, Nakata K, Mizugishi K, Aruga J, Mikoshiba K. 2001. Physical and functional interactions between Zic and Gli proteins. J Biol Chem 276:6889-92.

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CHAPTER 4: General Discussion

Heterotaxy is a disease of significant genetic heterogeneity. Clinically available gene panels now routinely offer targeted testing of close to 20 heterotaxy genes, while over 80 genes with involvement in L-R patterning have been described in animal systems (Rigler, et al., 2014).

As more genetic causes for heterotaxy have been identified, it has become increasingly evident that no single genetic abnormality is causative in a majority of patients. Diverse aneuploidies, chromosomal rearrangements, CNVs, and single gene mutations all explain a subset of disease, with each unique abnormality underlying only a small proportion of individual cases (Sutherland and Ware, 2009). Whereas our ability to detect disease-relevant genetic variation was once heavily restricted by technological limitations, state-of-the-art testing methods now provide significantly improved genomic coverage for considerably lower cost and effort. Ongoing technological refinements have helped to facilitate genome-wide genetic investigations as well as targeted sequence analyses of known or putative heterotaxy genes. In this dissertation, we have approached the study of heterotaxy from both of these levels, balancing comprehensive genome- wide gene discovery efforts with in-depth molecular and functional analysis of a known gene,

ZIC3, a primary genetic cause of X-linked heterotaxy, and a novel gene, PFKP.

4.1. PFKP AS A NOVEL CAUSE OF HETEROTAXY

We set out to identify novel genetic causes of heterotaxy by performing array-based CNV screens on a large cohort of 225 patients with diverse heart and situs defects. In agreement with high yields reported in previous analyses (Fakhro, et al., 2011; Rigler, et al., 2014), we identified

CNVs of clear or possible pathogenic significance in a large proportion (20.4%) of our cohort.

Detection of a subset of CNVs encompassing regions or genes already associated with

151 heterotaxy and CHD confirmed the ability of our assays to detect disease-relevant pathogenic variation and supported their capacity to identify novel heterotaxy genes. After filtering these and other CNVs not amenable to functional analysis, we were left with 35 rare CNVs of interest containing 165 unique genes suitable for candidate studies.

We selected the African clawed frog, Xenopus laevis, as the model system by which to screen promising gene candidates for relevance to L-R patterning pathways. The merits of

Xenopus for the study of laterality have been previously discussed both in this dissertation (see

Chapter 1.7) and elsewhere (Blum, et al., 2009). Most importantly, known laterality programs are conserved in Xenopus, with the ciliated gastrocoel roof plate (GRP) functioning analogously to L-R organizers (LROs) in mouse (the embryonic node), zebrafish (Kupffer’s vesicle, KV), and other vertebrate species (Schweickert, et al., 2007). Studies supporting earlier, ion-flux related laterality models have also derived significantly from work in Xenopus, providing additional pathways for functional interrogation (see Chapter 1.6).

Among our top heterotaxy candidates was the glycolytic enzyme, PFKP. From a genetic perspective, this gene was of interest as it was encompassed within a relatively compact (175kb) deletion not previously reported in control populations and clearly identified on both genotyping platforms. Functionally, although glycolytic factors have not yet been explicitly linked to L-R patterning, an association between maternal diabetes and increased incidence of heterotaxy in offspring of affected mothers has been long recognized (Morishima, et al., 1996; Slavotinek, et al., 1996; Splitt, et al., 1999; Loffredo, et al., 2001; Martinez-Frias, 2001; Lisowski, et al., 2010;

Lopez, et al., 2015). Xenopus-based morpholino knockdown experiments with assessments of organ situs and abnormal L-R molecular marker expression suggested PFKP to be a novel regulator of L-R patterning while ruling out involvement of PITRM1, the only other gene present

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in the CNV interval. A detailed hypothesis as to how PFKP might regulate L-R patterning has already been described elsewhere (see Chapter 2.5.2) and will not be repeated here.

Nevertheless, from a global perspective, the potential for PFKP to function as a regulator of L-R relevant ion flux (via H+-V-ATPase) places it in the context of one of the most fiercely debated questions in the field today: what are the earliest steps in L-R patterning? And to what extent are these mechanisms conserved among vertebrate species?

Such questions are largely framed by intense debate between supporters of the ion flux versus nodal flow models (see Chapter 1.6.3). Proponents of ion-flux believe that either (1) cleavage-stage ionic asymmetries are required for initiating events at the LRO, which serve as secondary amplification steps for the L-R patterning program, or that (2) these asymmetries operate independently of events at the LRO by feeding into the Nodal signaling cascade through one or more as-yet unidentified pathways (Vandenberg, et al., 2013). Opponents argue that the ability to establish the L-R axis is an intrinsic feature of the node and that no earlier cues are required (Beyer, et al., 2012; Blum, et al., 2014). As disruptions in both ion-flux and flow-related processes result in dysregulated Nodal signaling, it appears unlikely that these seemingly disparate models are entirely mutually exclusive.

In a recent review of completed work in Xenopus, Blum and colleagues (Blum, et al.,

2014), argue heavily in favor of the nodal flow model while providing an overview of competing evidence presented by the laboratory of Dr. Michael Levin (Vandenberg, et al., 2013). Blum et al. propose that serotonin is critical for L-R patterning not as a freely diffusible cleavage-stage determinant (as called for by the ion-flux model), but rather as a competence factor for Wnt- mediated specification of the presumptive GRP (Beyer, et al., 2012). Without serotonin, they argue, the GRP will not properly form and nodal flow will be interrupted. The authors

153 additionally hypothesize that other “early” determinants are likely to have unidentified roles at later LRO-stages and cite the gastric H+/K+ATPase, Atp4, as their representative example. Atp4 was the first published ion transporter with ties to L-R patterning and was originally identified through a series of pharmacological and genetic experiments in frog and chick (Levin, et al.,

2002). Levin et al. suggested a possible, ion-flux related role for Atp4 based on evidence that (1) atp4 mRNA is asymmetrically localized to the right-ventral blastomere in 4-cell stage Xenopus embryos, (2) Atp4 inhibition disrupts organ situs and left-sided expression of Nodal signaling components (Xnr-1, Pitx2, Lefty), and (3) Atp4 inhibition alters membrane potentials in streak- stage chick embryos. In their re-assessment of this data (Walentek, et al., 2012), Blum and colleagues noted that genetic or pharmacological knockdown of Atp4 in Xenopus yielded not only the previously noted organ situs and Nodal signaling defects, but also ciliary abnormalities and altered nodal flow. Additionally, dual roles for Atp4 in inducing Foxj1 expression in the presumptive GRP superficial mesoderm and in regulating polarization of GRP cilia were identified. At least in this case, disruption of a factor considered important for the ion flux model resulted in a number of abnormalities in systems relevant to nodal flow. Although it remains to be seen whether similar results will be obtained for other “early” factors (of which PFKP may be one), the onus, at least for the moment, lies with proponents of ion-flux to investigate what, if any, late-stage effects might be observed following experimental manipulation of putative early determinants (Blum, et al., 2014).

Future investigations into the L-R patterning role(s) of PFKP will need to take these timing issues into careful consideration to ensure that early versus late functions can be reliably distinguished. Relative to other model systems, methods for temporal control of gene expression in Xenopus are limited (Rankin, et al., 2011). Nevertheless, a number of existing protocols could

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be applied to the study of PFKP. First, the full-length human PFKP coding sequence could be subcloned into a pCSKA-GFP cytoskeletal actin promoter-driven Xenopus expression vector, which is expressed only after mid-blastula transition, Using this construct, PFKP overexpression would be restricted to stages relevant to “late” (nodal flow), but not “early” (ion-flux), L-R patterning processes {Pan, et al., 2006}. Second, transient transgenic embryos carrying a heat shock-inducible pfkp construct (pHSGFP-pfkp) could be generated (Wheeler, et al., 2008).

Induction after initial cleavages would again restrict dosage changes to “late” GRP-relevant stages. Finally pfkp MOs could be injected into 4-cell dorsal blastomeres, which are thought to give rise cells required for GRP specification, but not for ion-flux regulation (Blum, et al., 2009;

Vick, et al., 2009; Vandenberg and Levin, 2012). If these lineage specifications are strict and roles for pfkp in laterality are GRP-specific, then dorsally (but not ventrally) injected embryos should show L-R patterning defects similar to embryos injected at 2-cell stages. Ultimately, if pre-flow/ion-flux mechanisms are found to be important for establishing early asymmetries, it will be imperative to determine the extent to which these or other functionally similar processes are conserved across other vertebrate species.

4.2. ZIC3 MUTATIONS AS A CAUSE OF HETEROTAXY

Mutations in ZIC3 have long been recognized as a genetic cause of heterotaxy and CHD

(Gebbia, et al., 1997; Megarbane, et al., 2000; Ware, et al., 2004; Fritz, et al., 2005; Tzschach, et al., 2006; Chhin, et al., 2007; De Luca, et al., 2010; Wessels, et al., 2010; Chung, et al., 2011;

D'Alessandro, et al., 2011; Ma, et al., 2012; Cowan, et al., 2014). Sequencing efforts completed to date have collectively identified over twenty unique mutations in ZIC3 coding regions, the majority of which affect at least one of five centrally located zinc finger DNA binding domains

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(Stenson, et al., 2003; D'Alessandro, et al., 2013). Because these zinc finger sequences overlap critical nuclear import and export signals, mutations in DNA binding domains consistently alter both ZIC3 subcellular localization and target gene transactivation (Ware, et al., 2004; Bedard, et al., 2007; Hatayama, et al., 2008). These results have suggested a pathogenic mechanism whereby abnormal cytoplasmic localization of mutant ZIC3 is thought to preclude activation of as-yet unidentified developmentally critical gene targets. Mutations in flanking N- and C- terminal domains have been detected with much less frequency and are largely of uncertain functional consequence (Ware, et al., 2004; De Luca, et al., 2010; Wessels, et al., 2010; Cast, et al., 2012; D'Alessandro, et al., 2013).

In an effort to identify novel ZIC3 mutations and to better understand the pathogenesis of mutations detected throughout the gene, full ZIC3 coding regions and splice junctions were sequenced in 440 patients with heterotaxy or isolated CHD. Results from these sequencing efforts revealed a higher than anticipated mutation prevalence in sporadic heterotaxy, particularly among affected males. Our results additionally supported previously reported genotype- phenotype correlations between mutations in zinc finger regions and occurrence of laterality defects in patients with these mutations.

We performed in vitro functional testing on a total of 15 unique ZIC3 variants, including all 11 mutations identified in our heterotaxy cohort. Eight of these 11 mutations were novel to this study, including 5 in N- and C-terminal domains. Prior to this analysis, only 9 N-terminal variants had been previously reported, including 3 expansions of the N-terminal polyalanine tract

(Wessels, et al., 2010; D'Alessandro, et al., 2013). One of these, an expansion of only a single alanine residue (p.Ala53dup), was also detected in our cohort (Cowan, et al., 2014). Mutations of the polyalanine tract were of considerable interest to us as similar expansions are well recognized

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as causes of diverse genetic disease (Albrecht and Mundlos, 2005; Messaed and Rouleau, 2009).

A C-terminal variant detected in a position unique to ZIC3-A (p.Ala447Gly) was also particularly noteworthy as it represented the most C-terminal ZIC3 mutation yet reported. At present, only one other study (also completed by our group) has fully sequenced the alternative, ZIC3-B specific terminal exon (Bedard, et al., 2011). No mutations were identified in either analysis

(Bedard, et al., 2011; Cowan, et al., 2014).

Results from our functional studies confirmed previous findings that mutations affecting zinc finger regions significantly impair ZIC3 nuclear trafficking and transcriptional activation

(Ware, et al., 2004; Hatayama, et al., 2008). In contrast, N-terminal domain mutations, including those affecting the polyalanine tract, demonstrated little to no effect on these parameters. These results suggested one of two possibilities: either (1) mutations in non-zinc finger domains have no effect on ZIC3 function and are therefore unlikely to be disease causing, or (2) mutations in non-zinc finger domains impact distinct functions from mutations in DNA binding regions.

Although relatively fewer mutations have been detected in terminal domains, high levels of sequence conservation and low frequencies of polymorphisms in healthy controls indicate that there is potential for terminal mutations to have pathogenic significance. In what capacity might they be acting? As recent work has identified a role for ZIC3 in Wnt signaling, ostensibly via degradation of -catenin (Fujimi, et al., 2012; Ahmed, et al., 2013), one reasonable hypothesis is that mutations in terminal regions might affect the ability of ZIC3 to regulate appropriate levels of Wnt signals in L-R critical stages. This hypothesis appears to be unlikely, however, as missense mutations in N-terminal (Pro217Ala), but not in zinc finger domains (Trp255Gly,

Thr323Met), retain the ability of wild-type ZIC3 to inhibit a TOPFLASH Wnt reporter (Fujimi, et al., 2012). Similar results from a ZIC3 mutant that prematurely truncates before the five zinc

157 finger domains (p.E249X) support the idea that a role for ZIC3 in Wnt regulation is unlikely to be N-terminal related (Ahmed, et al., 2013). It is worth noting that C-terminal involvement cannot be explicitly ruled out as no C-terminal mutation has yet been explicitly tested; however, retention of Wnt-inhibition by a nearby ZF5 mutation (Lys405Glu), but not the previously described ZF1 (Trp255Gly) and ZF3 (Thr323Met) mutations, suggests that the Wnt-regulating potential of ZIC3 is more likely to lie further upstream.

If ZIC3 terminal domains do not contribute to subcellular localization, target gene activation, or Wnt signaling, in what capacity might they be operating? ZIC3 has been shown to be an enormously versatile gene (see Chapter 1.5.2, Chapter 4.3) and it is certainly plausible that intact terminal domains may be required for some as-yet-unidentified developmental function.

As novel roles for ZIC3 are identified, investigators should continue to assess both central and terminal mutations for functional significance. Likewise, re-assessment of terminal N- and C- terminal mutation frequency in affected versus control populations should be regularly undertaken to rule out occurrence of terminal mutations in the general population. Only in this way can definitive roles for terminal domains ultimately be established.

Polyalanine tract expansions beyond a certain threshold are thought to act as destabilizers of protein conformation, inducing misfolding of native structures and disruption of downstream protein functions (reviewed in Albrecht and Mundlos, 2005; Messaed and Rouleau, 2009).

Aberrant protein localization, degradation, aggregation, and interactions are direct molecular consequences and are presumed to impact downstream transcriptional activities and developmental functions. Both gain- and loss-of function are possible and increased severity is observed with larger expansion sizes. ZIC3 polyalanine tract expansions reported to date have included only one or two alanine residues (Wessels, et al., 2010; D'Alessandro, et al., 2013;

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Cowan, et al., 2014), contrasting rather sharply with the 10 alanine ZIC2 expansions reported to be causative for holoprosencephaly (Brown, et al., 1998; Brown, et al., 2001; Brown, et al.,

2005). The ZIC2 and ZIC3 polyalanine tracts share some fundamental differences, as the ZIC2 tract is C- rather than N-terminal and, at 15 residues, is longer than the 10 residue ZIC3 tract.

Functionally, the ZIC2 tract appears to suppress transactivation without effect on nuclear localization, as evidenced by increased activation and nuclear retention following deletion of the entire tract (Brown, et al., 2005). As our results indicate no change in either transactivation or subcellular localization with any ZIC3 polyalanine tract mutations, it is worth questioning (1) whether the differentially located ZIC3 and ZIC2 polyalanine tracts could possibly share similar functions in transcriptional repression and (2) whether a minimum number of residues must be affected or an expansion threshold must be reached before pathogenesis can occur. For the latter question, it is worth considering that an in-vitro generated 2-alanine expansion of the ZIC2 tract increased activation of an SV-40 luciferase reporter by 2.5-fold relative to wild-type ZIC2, indicating that even smaller expansions may have functional effects (Brown, et al., 2005). If they are ultimately determined to be functionally significant, the expansions identified in ZIC3 would be the shortest pathogenic expansions yet reported. Experiments both deleting and expanding the

ZIC3 polyalanine tract will be of considerable help in determining whether any functional similarities exist between it and its comparable sequence in ZIC2.

4.3. ZIC3 AND LEFT-RIGHT PATTERNING

Until recently, a precise developmental function for ZIC3 during L-R patterning had proven to be elusive. In a landmark finding, two reports, published simultaneously in 2013, demonstrated for the first time a role for Zic3 in mouse embryonic node formation (Jiang, et al.,

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2013; Sutherland, et al., 2013). Both studies (one of which was spearheaded by a former student in our lab) utilized a conditional loss-of-function approach to specifically delete Zic3 in tissues of suspected or known significance for laterality. Targeted deletion in the primitive streak and migrating mesoderm, but not in post-gastrulation node cells, yielded significant L-R patterning and cardiac defects accompanied by highly abnormal node ultrastructure and shortened nodal cilia. Cardiac defects were determined to have occurred secondarily to earlier defects in L-R patterning and not through cell autonomous mechanisms in the heart. The presence of ciliary abnormalities in conditional knockout mice places ZIC3 in the company of other heterotaxy genes with defined roles in ciliogenesis (see Chapter 1.6.1). Similar cilia length deficiencies have also been noted in the KV of zebrafish morphants, suggesting a possible conserved role for ZIC3 in cilia elongation (personal communication, Dr. Jeffrey Amack). By defining precise spatial and temporal requirements for ZIC3 in the LRO, these studies have begun to shed light on one of the major questions in L-R patterning and have re-concentrated efforts on determining specific mechanism(s) of action for ZIC3 in laterality.

Some tantalizing clues are provided by the complex phenotypes displayed by ZIC3 null mice, including a variety of abnormalities stemming from dysregulated gastrulation and neurulation (Purandare, et al., 2002; Ware, et al., 2006b). Progression through gastrulation, elongation of the anterior-posterior body axis, and closure of the neural tube all depend on convergent extension (C.E.), a type of cell movement characterized by elongation of one embryonic axis and simultaneous narrowing of a second. Using morpholino-based approaches, our lab has previously demonstrated a requirement for ZIC3 in regulating C.E. movements in frog and zebrafish (Cast, et al., 2012): in addition to organ situs and molecular marker abnormalities indicative of disrupted L-R patterning, morphant embryos in this study displayed

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defects in blastopore closure, notochord elongation, anterior-posterior (A-P) extension, and somite and neural plate development - all indicators of faulty C.E. As these phenotypes indicate a critical role for ZIC3 in C.E. at time points relevant to node formation, it is tempting to speculate that Zic3 functions in both processes may be related.

C.E. is regulated by the same planar cell polarity (PCP) pathways that govern basal body posteriorization and axonemal tilting of nodal cilia (Ybot-Gonzalez, et al., 2007; Maisonneuve, et al., 2009; Antic, et al., 2010; Borovina, et al., 2010; Hashimoto, et al., 2010; Song, et al., 2010;

Wallingford and Mitchell, 2011; Wang, et al., 2011; Kim, et al., 2012). Accumulating evidence has also linked PCP proteins to ciliogenesis, adding an additional level of molecular genetic complexity to cilia assembly programs (reviewed in Wallingford and Mitchell, 2011).

Preliminary studies completed in our lab (unpublished, Mardi Sutherland) have identified a number of differentially expressed PCP genes in node cells from Zic3 null mice, suggesting that

Zic3 may play a role in regulating PCP during L-R patterning. Both up-and down-regulated genes were identified, including Wnt5a, a recognized PCP pathway activator (Gray, et al., 2011), and Dvl1, a core PCP protein required for cilia posteriorization (Hashimoto, et al., 2010). Future experiments focusing on the structural and molecular consequences of ZIC3 knockdown on PCP- regulated processes will be required to determine whether Zic3 acts primarily as a transcriptional activator or repressor in this developmental context.

4.4. THE FUTURE OF GENETIC TESTING FOR HETEROTAXY

In this dissertation, we have described results from comprehensive genetic analyses of two large heterotaxy cohorts. These analyses, ranging from full gene sequencing to comprehensive genome-wide CNV screens, have expanded our understanding of the diverse

161 genetic causes of heterotaxy while identifying a strong list of candidate genes for future study.

Recent years in particular have seen significant advancements in the resolution and affordability of genome-wide approaches to the study of genetic disease, with the successful application of microarray technologies to the study of heterotaxy being only the most relevant example

(Fakhro, et al., 2011; Rigler, et al., 2014).

Emerging massively parallel next-generation sequencing (NGS) approaches have begun to revolutionize genomic approaches to novel gene discovery and patient diagnosis. Improved genomic coverage and faster turnaround times relative to traditional capillary-based sequencing methods (Shendure and Ji, 2008; Mardis, 2011) have spurred widespread implementation of these technologies in both clinical and research settings (Bamshad, et al., 2011; Schuler, et al.,

2013; Atwal, et al., 2014; Iglesias, et al., 2014; Levenson, 2014). Like microarray technologies, the major strength of NGS has been its ability to detect genetic variation without a priori knowledge of causative disease genes. This capability has not only aided gene discovery efforts, but also assisted diagnostic evaluation of patients with genetic diseases of uncertain etiology or complex conditions stemming from mutations in large numbers of causative loci. In large part, the versatility of these sequencing methods derive from their scalability - both to large-scale whole genome (WGS) or whole exome (WES) analyses and to targeted gene sequencing. On the genome-wide level, NGS has already been applied to the study of congenital heart disease

(reviewed in Dorn, et al., 2014) and to heterotaxy (Tariq, et al., 2011). The latter study, in which we which identified SHROOM3 as a novel heterotaxy gene (Tariq, et al., 2011), clearly demonstrates the enormous potential of these technologies for the study of patients with laterality disorders. Ongoing application of NGS technologies to these patient populations will no doubt continue to provide novel insights into known and as-yet unidentified L-R patterning pathways.

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Despite the undeniable promise of these technologies for the study of heterotaxy, their rapid development and widespread implementation has underscored a number of increasingly relevant concerns, particularly with respect to interpretation and reporting of variants of uncertain significance. Our ability to interpret detected variation is often limited by a lack of understanding of functional genomic context, particularly when identified variants are found in non-coding regions. From a data management perspective, the sheer volume of identified variation can also be daunting. WES identifies, on average, 12,000 unique coding variants per exome sequenced (Ng, et al., 2008) and potentially pathogenic variants are observed even in apparently healthy individuals (Li, et al., 2010; Marth, et al., 2011; Tennessen, et al., 2012).

Return of incidental findings, availability of insurance coverage, and cost effectiveness relative to existing multi-gene panels have all been raised as additional clinical concerns (Kaye, et al.,

2010; Atwal, et al., 2014). At a minimum, it has become an expectation that reporting laboratories will perform thorough literature reviews for all variants of potential clinical relevance and properly classify each as having known or uncertain significance. Despite these limitations, it is anticipated that the already widespread application of NGS will lead to improvements to existing technologies and to databases of normal and disease-related genetic variation. These refinements will no doubt aid in decreasing overall costs and improving accuracy of variant interpretation.

4.5. FINAL REMARKS

The rapid growth of genome-wide testing technologies has facilitated unprecedented gains in genetic coverage for historically low time and monetary investments. As we enter the next phase of heterotaxy genetic study our greatest challenge will lie in parsing and interpreting

163 the wealth of anticipated genomic data, particularly as the field advances to consider contributions of susceptibility factors and common variation to genetic disease as well as multifactorial and oligogenic disease models. Continued implementation of available testing methods will undoubtedly lead to improvements to existing technologies and databases of normal and disease-related variation. As a community, these ongoing efforts will ensure that our collective push towards novel discovery does not outpace our ability to accurately interpret disease-relevant findings and integrate them within the larger context of known L-R patterning paradigms.

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