Williams Syndrome Transcription Factor is critical for neural crest cell function in Xenopus laevis

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Williams Syndrome Transcription Factor is critical for neural crest cell function in Xenopus laevis Chris Barnett a, Oya Yazgan a, Hui-Ching Kuo a, Sreepurna Malakar a, Trevor Thomas a, Amanda Fitzgerald a, William Harbour a, Jonathan J. Henry b, Jocelyn E. Krebs a,* a b

Department of Biological Sciences, University of Alaska Anchorage, 3101 Science Circle, Anchorage, AK 99508, USA Department of Cell and Developmental Biology, University of Illinois, 601 S. Goodwin Avenue, Urbana, IL 61801, USA

A R T I C L E I N F O

A B S T R A C T

Article history:

Williams Syndrome Transcription Factor (WSTF) is one of 25 haplodeficient genes in

Received 6 January 2012

patients with the complex developmental disorder Williams Syndrome (WS). WS results

Received in revised form

in visual/spatial processing defects, cognitive impairment, unique behavioral phenotypes,

31 May 2012

characteristic ‘‘elfin’’ facial features, low muscle tone and heart defects. WSTF exists in sev-

Accepted 1 June 2012

eral chromatin remodeling complexes and has roles in transcription, replication, and

Available online 9 June 2012

repair. Chromatin remodeling is essential during embryogenesis, but WSTF’s role in vertebrate development is poorly characterized. To investigate the developmental role of WSTF,

Keywords: WSTF Neural crest Xenopus Williams Syndrome BAZ1b Chromatin remodeler

we knocked down WSTF in Xenopus laevis embryos using a morpholino that targets WSTF mRNA. BMP4 shows markedly increased and spatially aberrant expression in WSTF-deficient embryos, while SHH, MRF4, PAX2, EPHA4 and SOX2 expression are severely reduced, coupled with defects in a number of developing embryonic structures and organs. WSTFdeficient embryos display defects in anterior neural development. Induction of the neural crest, measured by expression of the neural crest-specific genes SNAIL and SLUG, is unaffected by WSTF depletion. However, at subsequent stages WSTF knockdown results in a severe defect in neural crest migration and/or maintenance. Consistent with a maintenance defect, WSTF knockdowns display a specific pattern of increased apoptosis at the tailbud stage in regions corresponding to the path of cranial neural crest migration. Our work is the first to describe a role for WSTF in proper neural crest function, and suggests that neural crest defects resulting from WSTF haploinsufficiency may be a major contributor to the pathoembryology of WS. Ó 2012 Elsevier Ireland Ltd. All rights reserved.

1.

Introduction

The development of a multicellular organism is a period of extensive transcriptional regulation, which is highly dependent on chromatin remodeling. During development, specialized cells arise from the gradual loss of pluripotency in the progeny of embryonic stem cells. Recent studies reveal the

importance of dynamically regulated chromatin structure in the maintenance of pluripotency and in cell differentiation (Delgado-Olguin and Recillas-Targa, 2011). WSTF (Williams Syndrome Transcription Factor), encoded by the gene WSCR9/BAZ1B, is one of about 25–30 contiguous genes that are haplodeficient in individuals with Williams Syndrome (WS). WS is an autosomal dominant disorder

* Corresponding author. Tel.: +1 907 786 1556; fax: +1 907 786 4607. E-mail addresses: [email protected] (C. Barnett), [email protected] (O. Yazgan), [email protected] (S. Malakar), [email protected] (T. Thomas), [email protected] (A. Fitzgerald), [email protected] (W. Harbour), j-henry4@illinois. edu (J.J. Henry), [email protected] (J.E. Krebs). 0925-4773/$ - see front matter Ó 2012 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.mod.2012.06.001

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occurring in 1 in 8000 live births and is the result of a 1.5 megabase deletion on chromosome 7 (Lu et al., 1998). Individuals with WS exhibit characteristic malformations of craniofacial, heart and neural structures, infantile hypercalcaemia, developmental delays, and complex and distinctive cognitive and behavioral profiles. The contributions of most of the deleted genes to the clinical outcomes of WS patients are not clear, though a handful of genes have been assigned tentative roles (reviewed in Osborne, 2010). Evidence from mouse models suggests that WSTF may play a critical role in heart and craniofacial development, and WSTF hemizygosity may also explain the hypercalcaemia observed in WS patients (Ashe et al., 2008; Yoshimura et al., 2009). WSTF is a subunit of three well-characterized ATPdependent chromatin remodelers: WINAC (WSTF including the nucleosome assembly complex), WICH (WSTF-ISWI chromatin remodeling complex) and B-WICH. WSTF is a 170 kDa protein in the BAZ/WAL family, characterized by six sequential motifs, including motifs implicated in chromatin binding. WSTF has roles in DNA repair, replication, transcriptional activation and repression, and also possesses histone H2A kinase activity (recently reviewed in Barnett and Krebs, 2011). In humans, WSTF transcripts are detected in all sampled adult tissues (heart, brain, placenta, skeletal muscle, and ovary) and four fetal tissues (brain, lung, kidney, and liver) in non-WS individuals (Lu et al., 1998). WSTF mRNA is detected in Xenopus laevis oocytes and is ubiquitous until early neurulation, when WSTF expression becomes confined to the neural ectoderm and closing neural tube (Cus et al. 2006). As neural tissue continues to differentiate into the early tadpole stage, WSTF mRNA localizes to specific neural structures and cells including the neural tube, optic cup, anterior brain, and migrating cranial neural crest cells (Cus et al., 2006). WSTF also displays strong expression in the branchial (pharyngeal) apparatus, a primordium for a number of organs and facial structures, such as the thyroid, aorta, the maxillary and mandibular bones, ligaments, and nerves as well as auditory structures like the auditory tube, bones of the middle ear and the auricles. This branchial arch expression appears to be limited to a population of cells located deep to the branchial arch ectoderm and is likely the contribution of cranial neural crest cells that originate from a space just dorsal to the neural tube; a number of these cells migrate to the branchial arches and play a crucial role in the proper development of this important primordial structure (Noden and Schneider, 2006). The neural crest is a unique group of embryonic cells found only in vertebrates (Gans and Northcutt, 1983). The neural crest originates from the ectoderm of the neural plate border that undergoes an epithelial-to-mesenchymal transition, giving rise to neural crest cells during development. This transition requires the orchestrated activities of a network of transcription factors that gradually lead to this transition (reviewed in Sauka-Spengler and Bronner-Fraser, 2008). Neural crest cells possess migratory ability as well as the ability to undergo extensive cellular reprogramming, which results in an increase in differentiation potential. The pluripotency of neural crest cells allows them to contribute to a number of developing tissues, including the central and peripheral nervous system, as well as many non-neural structures such as

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bone, cartilage, connective tissue, skin (melanocytes), and smooth muscle. The migration and maintenance of the neural crest cells is guided by morphogens, signaling molecules that are secreted from surrounding tissues. Bone morphogenic protein 4 (Bmp4) is a key developmental morphogen, and both inhibition and activation of the BMP4 gene is necessary for proper induction and maintenance of neural crest (Steventon et al., 2009). Additionally, both overexpression and inhibition of BMP4 alters facial development in chick, suggesting accurate timing and expression of BMP4 is crucial to proper neural crest function (Creuzet et al., 2002; Ruhin et al., 2003; Wawersik et al., 2005). Sonic hedgehog (Shh) is another morphogen that along with Bmp4 is critical for the proper patterning of the neural tube during embryogenesis (sometimes playing antagonistic roles) and is also important in craniofacial development of chick, human and mouse (Briscoe et al., 1999; Campuzano and Modolell, 1992; Owens et al., 2005; Roelink et al., 1995). Shh acts as an anti-apoptotic signaling molecule for neural crest cells (Ahlgren et al., 2002). Neural crest cells migrate in specific segmented regions between the overlying ectoderm and underlying mesoderm. The pathways of cranial neural crest cells are highly conserved in vertebrate development and proper axial patterning and organization of the region is critical for proper formation and migration of cranial neural crest (reviewed in Basch and Bronner-Fraser, 2006). Migration occurs from the forebrain and rhombomeres of the hindbrain to regions in the head and to the branchial arches (Noden and Schneider, 2006). The contributions of neural crest cells are both unique to and critical for the proper development of all vertebrates. Little is known about the role of chromatin remodeling in the formation of the neural crest and migrating neural crest cells. One recent study has shown that Chd7 (Chromodomain helicase DNA-binding domain member 7), an ATP-dependent chromatin remodeler, is essential for early neural crest formation in Xenopus embryos (Bajpai et al., 2010). In humans, heterozygous mutation of CHD7 leads to the congenital developmental disorder CHARGE Syndrome, characterized by specific craniofacial features, neural defects and malformations in the eyes, ears and heart (Pagon et al., 1981). Within the PBAF complex (Polybromo- and BRG1-Associated Factor-containing complex), Chd7 has been shown to bind in vivo to promoter-distal elements of the neural crest specifier SOX9 and the neural crest gene Twist in human neural crest-like cells (Bajpai et al., 2010). A number of the clinical features of WS involve structures that are derived from the neural crest, including craniofacial structures, heart and neural tissue. Given this observation, and the expression of WSTF in neural crest in X. laevis, we sought to determine the effects of WSTF knockdown in Xenopus embryos, and to particularly address the effects of WSTF knockdown in neural crest formation. We show specific defects in neural crest cell migration and/or maintenance in WSTF knockdown Xenopus embryos and propose a model in which misregulation of BMP4 and SHH expression in WSTF knockdowns results in the failure of normal neural crest contributions during development. Our data suggest that a number of the symptoms present in WS patients could be due to

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WSTF haploinsufficiency and the downstream effects of this multifunctional chromatin remodeler subunit.

2.

Results

2.1. WSTF Morphant embryos exhibit severe defects in neural development To determine the function of WSTF in early development, an anti-WSTF morpholino (MO) was injected into fertilized X. laevis eggs at the one-cell stage to inhibit the translation of endogenous WSTF mRNA. Eggs were injected with 42 ng of either anti-WSTF MO, or with a control MO that is the inverse of the anti-WSTF MO sequence (INV-MO). The inverse MO-injected embryos are indistinguishable from uninjected controls. Anti-WTSF MO injection results in an average reduction to 50–60% of endogenous WSTF levels, comparable to the hemizygous state in WS patients (Howald et al., 2006; Merla et al., 2006). Complete knockdown in early stages is not achievable due to the presence of high levels of maternally-deposited WSTF protein present in the developing embryos. Knockdown of WSTF protein levels was confirmed by Western blot (Supplementary Fig. 1). WSTF knockdown embryos progress normally through the early stages of development, at least at the gross morphological level. However, severe abnormalities become evident once embryos reach about stage 35, late tailbud stage (Fig. 1A and B) (Nieuwkoop and Faber, 1967). WSTF knockdown embryos display abnormal craniofacial development, including a deformed cement gland, an unusually pronounced indentation above the cement gland (in the region that will develop into the olfactory organs, primary mouth and anterior pituitary; Dickinson and Sive, 2007), and a

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protruding/misshapen forehead. In addition, the developing eyes are undersized and irregularly shaped, and the embryos exhibit exopthalmia (protruding eyes). Finally, the WSTF knockdown embryos show abnormal A–P axis development and particularly strong axial curvatures. By stage 41 the developmental abnormalities are more severe and the embryos die around late tadpole stage, about stage 45. Injections were performed in numerous independent batches of hundreds of embryos, with each injection resulting in 100% penetrance of this gross morphological phenotype (for example, in a typical experiment 333/355 embryos, or 94%, injected with anti-WSTF MO exhibit the strong WSTF phenotype, while 22/355, or 6%, show a milder phenotype mostly characterized by eye defects). Injection of the INV-MO typically resulted in non-specific developmental defects in <10% of embryos (for example, in a typical experiment 9/124, or 7%, embryos exhibit various defects). To obtain a more detailed understanding of the phenotypes of the WSTF knockdown embryos, we performed histological analysis to further characterize the anterior neural defects. Transverse sections through the anterior region of the embryos reveal details of the defects in eye and brain development in the WSTF knockdown embryos. Eye development is profoundly disrupted (Fig. 1C and D). The optic cups are generally shallow and poorly shaped, though in most (but not all) cases the pigmented retinal epithelium is present. The multiple layers of the neural retina completely fail to differentiate and are typically present as a solid mass of undifferentiated cells. The optic vesicle often retains a substantial attachment with the forebrain, without a clear optic nerve. Furthermore, the lens is occasionally absent, or present merely as a minimally differentiated thickening within the surface ectoderm, though in a few cases some primary lens fibers are detectable. The lenses invariably fail to detach

Fig. 1 – WSTF knockdowns exhibit neural defects and severely impaired eye development. A–B: Embryos (anterior to the left) injected at the one-cell stage with either WSTF-inverse control MO (top of each panel) or WSTF MO (bottom) at stages 37 (A) and 41 (B). The WSTF MO injected embryos exhibit reduction and malformation of neural structures, deformed eyes, and axial defects. Asterisks indicate the pronounced concavity at the site of the presumptive primary mouth. C-F: Stained histological sections of control morpholino-injected embryos (C, E) and of WSTF knockdown embryos (D, F), all injected at the one-cell stage. WSTF knockdowns display perturbed lens and retinal cell differentiation (D) and profound disorganization of neural tissues (F). e: eye; cg: cement gland; l: lens; r: retina; pe: pigmented retinal epithelium; nt: neural tube; nc: notochord. Scale bars: 1 mm (A and B) and 100 lm (C, D, E & F).

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from the ectoderm, and instead remain tightly associated with this layer. Many lenses protrude outward from the surface ectoderm. Organization of tissues within the developing brain of WSTF knockdown embryos is greatly affected as well (Fig. 1E and F). Neural tissue in the developing brain is poorly formed and differentiated. There appears to be a lack of cell proliferation and absence of a clearly defined proliferative (ventricular) zone. In some cases it appears that there is incomplete fusion of the neural tube. The prechordal mesoderm and notochord are frequently malformed, possibly enlarged in diameter and poorly differentiated. The whole mounts and sections of embryos in which WSTF has been globally knocked down (i.e. morpholino injected at the one-cell stage) clearly show that neural development is perturbed, and neural tissues are highly disorganized. It also appears that the amount of neural tissue is reduced in these embryos, at least in the anterior. This apparent reduction of neural tissue formation was confirmed by analyzing the expression of the neural-specific marker Neural Cell Adhesion Molecule (NCAM) by whole mount in situ hybridization (Fig. 2; representative embryos shown from a sample of 8–10 embryos for each stage). In addition to the embryos in which WSTF expression has been globally knocked down,

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we also examined embryos that were injected with antiWSTF MO into one cell of an embryo at the two-cell stage of development (Fig. 2A–C). This results in an embryo in which the MO affects the embryo unilaterally and thus only exhibits the knockdown phenotype on the injected side, while the other side serves as the control. The injected side is traced by visualizing the fluorescently tagged MO (Fig. 2A). NCAM expression is reduced in the injected side of a unilaterally-injected two-cell stage embryo post-neurulation at stage 21, particularly in the anterior region and the eye (Fig. 2B and C). NCAM expression is also reduced in embryos where WSTF has been globally knocked down. WSTF globalknockdown embryos at a late tailbud stage (stage 35) show a severe reduction of NCAM expression compared to INVMO injected controls, especially in the brain and branchial arches (Fig. 2D and E). These data indicate that WSTF is important for normal neural tissue development.

2.2. BMP4, SHH and MRF4 genes are misregulated in WSTF knockdown embryos To further investigate the effects of WSTF knockdown on neural development, we examined two genes known to regulate neural proliferation and differentiation during embryonic

Fig. 2 – WSTF function is required for neural tissue formation in vivo. A: Two-cell stage embryos (dorsal view, anterior at top) were injected with carboxyfluorescein-labeled WSTF MO unilaterally into a single blastomere; fluorescence indicates the injected side. B-C: NCAM expression analyzed by whole mount in situ hybridization at late neurulation (stage 21; B: dorsal view, anterior at top; C: anterior view, dorsal at top). NCAM is normally expressed in the eyes and neural ectoderm. NCAM signal is diminished in the anterior on the WSTF MO-injected side. Asterisk denotes region of greatest loss of NCAM signal. C: Anterior view of embryo in (B), highlighting anterior reduction of NCAM expression in the eye rudiments and neural ectoderm on the injected side. Asterisk denotes loss of NCAM signal. D-E: NCAM expression analyzed by whole mount in situ hybridization at late tailbud stage (stage 35; side views, anterior to left). D: One-cell stage embryos were injected with WSTF-inverse MO (INV) and display normal NCAM expression in neural ectoderm, eye (e) and branchial arches (ba). E: One-cell stage embryos injected with WSTF MO (MO) display reduced NCAM expression at late tailbud stage. Bracket marked by asterisk indicates loss of NCAM staining in branchial arches in WSTF-MO injected embryos (E). Scale bars: 0.5 mm (A–C) and 1 mm (D and E).

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development. Bmp4 is a signaling molecule from the TGFb/ BMP family of proteins known for neural patterning, cell differentiation and cell death (reviewed in Hogan, 1996). The expression domain of BMP4 is significantly expanded on the injected side of a unilaterally-injected embryo during late neurulation (stage 17; Fig. 3A; representative embryos shown from a sample of 12 embryos for each stage). While normal

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BMP4 expression is displayed in embryos globally injected with inverse morpholino, in global WSTF knockdowns, BMP4 expression is markedly increased in late tailbud stage embryos (stage 40; Fig. 3B), as well as in earlier tailbud embryos (stage 37) as measured both by in situ hybridization and RTqPCR of whole embryos (Fig. 3E and data not shown). Strikingly, BMP4 expression is spatially misregulated. BMP4

Fig. 3 – SHH, BMP4 and MRF4 are misexpressed in WSTF knockdown embryos. A: BMP4 expression in the neural plate analyzed by whole mount in situ hybridization at late neurulation (stage 17; dorsal view, anterior at top) in embryos injected with WSTF morpholino at the two-cell stage. BMP4 expression is expanded beyond its normal domain on the WSTF knockdown side (left) compared to the uninjected control side (right). B: Whole mount in situ hybridization displaying normal expression pattern of BMP4 compared to global WSTF knockdown embryos at late tailbud stage (stage 40; side view, anterior to left). One-cell stage embryos were injected with either WSTF MO (bottom) or WSTF-inverse MO (INV, top). Inverse MO globally injected embryos display normal BMP4 expression in the hindbrain, developing heart, and faintly in the otic vesicle. Global WSTF knockdowns display decreased BMP4 anterior expression as well as aberrant expression in dorsal tissues. C: Whole mount in situ hybridization displaying normal expression of pattern SHH compared to global WSTF knockdowns at late tailbud stage (stage 36–37; side view, anterior to left). One-cell stage embryos were injected with either WSTF MO (bottom) or WSTF-inverse MO (top). Globally injected inverse MO embryos display SHH expression in the head and notochord. Global WSTF knockdown embryos display decreased SHH expression throughout. D: Global WSTF knockdown embryo (right) displaying cyclopia/holoprosencephaly compared to globally injected inverse MO embryo (left) at stage 45 (dorsal views of heads, anterior at top). Arrows indicate location of eye(s). One-cell stage embryos were injected with either WSTF MO or WSTF-inverse MO. E: Real time RTPCR results for BMP4 and SHH graphed as a percentage of normal expression. BMP4 (stage 37) is significantly over-expressed and SHH expression (stage 15) is significantly reduced in WSTF knockdown embryos injected at the one-cell stage. Data are the average of 3–4 independent experiments and standard errors are shown. F: MRF4 expression in a stage 15 embryo (dorsal view, anterior at top) injected with WSTF MO at the two-cell stage. The injected side (left) displays a significant decrease in normal MRF4 expression in the developing somites. Scale bars: 0.5 mm (A and F) and 1 mm (B–D).

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exhibits reduced expression within its normal expression domain, specifically in anterior structures like the heart and hindbrain (Martinez-Barbera et al., 1997). In addition, WSTF morphants show a marked increase of BMP4 expression within tissues around the dorsal midline that include notochord and somitic mesoderm, tissues that do not normally express BMP4 (Fig. 3B and Supplementary Fig. 2). Shh is known for its role in midline signaling and the splitting of bilateral eye fields, and is essential for survival of a number of neural cell types during embryogenesis (Chiang et al., 1996; Prykhozhij, 2010). SHH transcripts are significantly decreased in both the anterior regions and neural tube in global WSTF knockdown embryos compared to controls (Fig. 3C). Mutations in SHH are linked to holoprosencephaly, the failure or incomplete division of the two hemispheres of the forebrain, and cyclopia, the failure or incomplete separation of the eye fields (Dubourg et al., 2007). Consistent with the reduced SHH expression we observe, WSTF global knockdown embryos that survive well into the tadpole stage exhibit a severe anterior phenotype characterized by cyclopia (Fig. 3D). Misregulation of SHH (at stage 15) and BMP4 (at stage 37) was also confirmed by real time RT-PCR (Fig. 3E). SHH is expressed in the notochord, a mesoderm-derived tissue that defines the axis of all chordates (Yamada et al., 1993). To further investigate the effect of WSTF depletion on mesoderm development we assayed the expression of MRF4, a marker for somites, also a mesoderm-derived tissue. Injection of anti-WSTF MO into one cell of two-cell stage embryos, results in embryos in which only half of the embryo is affected. MRF4 expression in WSTF unilateral-knockdown embryos is reduced on the injected side during neurulation (stage 15; Fig. 3F). These results indicate that in WSTF knockdowns, both ectoderm- and mesoderm-derived tissues show perturbed gene expression, including overexpression and spatially aberrant expression of BMP4, and reduced expression of mesoderm-derived tissue markers SHH and MRF4.

2.3. WSTF knockdown embryos exhibit perturbed PAX2, EPHA4, and SOX2 expression While the reduced expression of NCAM and the sections of WSTF knockdown embryos clearly indicate a reduction in neural tissue in these embryos, we wished to further examine the differentiation of anterior neural tissues. The Paired box 2 (PAX2) gene encodes a Pax family transcription factor with important roles in vertebrate organogenesis (Carroll and Vize, 1999; Dressler et al., 1990). PAX2 expression is crucial for development of the midbrain hindbrain boundary (MHB) in vertebrates, which is required for neuronal differentiation and patterning of the midbrain and anterior hindbrain (Li Song and Joyner, 2000). We examined PAX2 expression via whole mount in situ hybridization to investigate midbrain and cerebellum development in our WSTF knockdown embryos. Fertilized embryos were injected with anti-WSTF MO at the one cell stage, resulting in global knockdown of WSTF. WSTF knockdown embryos at tailbud stage display an almost complete loss of PAX2 expression within the hindbrain and a severe reduction at the MHB

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when compared to controls (Fig. 4A–C; representative embryos shown from a sample of 5 control and 7 knockdown embryos). Additionally, at this stage, expression of PAX2 is vital to the formation of the pronephros, the nascent yet functional embryonic kidney (Heller and Brandli, 1997). PAX2 signal is also reduced when compared to controls, particularly at the anterior end of this structure, suggesting that normal kidney development may be perturbed in our WSTF morphant embryos (Fig. 4A and B). Eph receptors are a family of receptor tyrosine kinases that bind a family of ephrin ligands involved in numerous developmental processes. EPHA4 is expressed in a number of tissues during Xenopus development including the forebrain, hindbrain, pronepheros, and the olfactory and otic placodes (Park et al., 2004; Winning and Sargent, 1994; Xu et al., 1995). In Xenopus, EPHA4 is involved in neural tissue development in the forebrain and the MHB, and plays a crucial role in neural crest cell migration (Smith et al., 1997; Xu et al., 1995). We examined EPHA4 expression by whole mount in situ hybridization to further investigate its role in neural development in our WSTF knockdown embryos. Fertilized embryos were injected with anti-WSTF MO at the one cell stage, resulting in global knockdown of WSTF in the developing embryos. Consistent with PAX2 expression, knockdown embryos at the tailbud stage display a severe reduction of EPHA4 expression within the forebrain and MHB when compared to controls, further indicating failure of neural tissue differentiation at this stage (Fig. 4). Normal expression of EPHA4 in rhombomeres 3 and 5 is also reduced in WSTF knockdown embryos (Fig. 4D–F; representative embryos shown from a sample of 4 control and 8 knockdown embryos) (Smith et al., 1997). Rhombomere formation is crucial for the segmentation and proper migration of neural crest cells (Smith et al., 1997). This reduction in EPHA4 could result in underdeveloped rhombomeres and improper neural crest migration in our WSTF morphant embryos. Additionally, as with PAX2, expression of EPHA4 in the pronephros may be reduced, and the pronephros is smaller and deformed compared to the controls (Fig. 4E). Sox2 is a transcription factor well known for its role in maintaining the pluripotency of embryonic stem cells (Heng et al., 2010). In addition, Sox2 in combination with different binding partners plays numerous roles in embryonic development (Archer et al., 2011; Kondoh and Kamachi, 2010), including development of the CNS and the retina and lens of the eye. In the developing brain, Sox2 has dual roles in maintaining proliferative neural stem cells as well as being required for specific neuronal differentiation (Agathocleous et al., 2009; Cavallaro et al., 2008; Pevny and Nicolis, 2010). As above, we examined SOX2 expression by whole mount in situ hybridization in our WSTF knockdown embryos. In this case, fertilized embryos were injected with anti-WSTF MO at the two cell stage, resulting in unilateral knockdown of WSTF in one side of the developing embryos. Again consistent with the other neural markers, the MO-injected side at the early tailbud stage displays a severe reduction of SOX2 expression throughout the brain, eye, otic vesicle, lateral line placodes, and branchial arches when compared to the uninjected control side (Fig. 4 G–J; representative embryos shown from a sample of 4–6 embryos for each stage).

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Fig. 4 – PAX2, EPHA4, and SOX2 expression are reduced in WSTF knockdown embryos. A: PAX2 expression in the midbrain hindbrain boundary (MHB), the otic vesicle, optic stalk and the pronephros analyzed by whole mount in situ hybridization in control embryo at tailbud stage (Stage 33; lateral view, anterior at left) B: Tailbud stage embryo injected with WSTF morpholino at the one-cell stage. PAX2 expression is reduced in all expression domains. C: Side-by-side dorsal view of control embryo (left) with WSTF knockdown embryo (right) at tailbud stage, showing reduced expression of PAX2 in the MHB and the otic vesicle of WSTF knockdown embryo compared to control (Stage 33; dorsal view, anterior at top). D: EPHA4 expression in the forebrain, hindbrain, rhombomeres 3 and 5 and in the pronephros analyzed by whole mount in situ hybridization in control embryo at tailbud stage (Stage 33; lateral view, anterior at left) E: Tailbud stage embryo injected with WSTF morpholino at the one-cell stage. EPHA4 expression is reduced in all expression domains. F: Side-by-side dorsal view of control embryo (left) with WSTF knockdown embryo (right) at tailbud stage showing reduced expression of EPHA4 in the forebrain, hindbrain and rhombomeres 3 and 5 of WSTF knockdown embryo compared to control (stage 33; dorsal view, anterior at top). G-J: Tailbud stage embryos injected with WSTF morpholino at the two-cell stage. G: SOX2 expression in the brain, eye, otic vesicle, lateral line placodes and branchial arches analyzed by whole mount in situ hybridization on the uninjected side at tailbud stage (Stage 32; lateral view, right side of embryo). H: SOX2 expression is reduced in all expression domains on the injected side (left side of embryo). I and J: Dorsal views of two embryos injected at the two-cell stage (WSTF MO injected into the left side in both cases). Embryo in I is at stage 28, embryo in J is stage 32. Both show reduced expression of SOX2 in all expression domains on the injected side. STD: Standard control morpholino; MO: WSTF morpholino; os: optic stalk; br: brain; pd: pronephric duct; hb: hindbrain; pn: pronephros, MHB: mid-hindbrain boundary; ov: otic vesicle; r3, r5: rhombomeres 3 and 5; op: olfactory placode; cp: cranial placodes [otic and lateral line]; ba: branchial arches; e: eye. Asterisks indicate areas of reduced expression relative to controls. Scale bars: 0.5 mm (A, B, D, E, G, H), 0.1 mm (C, F, I, J).

2.4. WSTF is crucial for proper neural crest cell function in vivo WSTF mRNA expression is detected in both the migratory neural crest as well as branchial arches during Xenopus embryogenesis (Cus et al., 2006). This expression, and the reduced expression of SHH and EPHA4, genes known to play roles in neural crest function, suggest that WSTF knockdown might impact the neural crest. In the anterior, neural crest cells originate from just above the neural tube and migrate

to the branchial arches, after which they serve as a critical component for the formation of a number of subsequent anterior structures that originate from the branchial arches. SNAIL and SLUG are two members of the Snail family that function in the epithelial-to-mesenchymal transition cascade. SLUG is expressed in the premigratory neural crest cells in both chick and X. laevis (Mayor et al., 1995, 1999; Nieto et al., 1994; Sefton et al., 1998). In Xenopus, SNAIL is expressed in ectoderm, premigratory neural crest, and the branchial arches (Mayor et al., 1993). In many vertebrates, SNAIL and SLUG

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play important and seemingly redundant roles in the initiation of neural crest formation (Aybar et al., 2003). However, in Xenopus, SNAIL appears to be an initiator of the neural crest genetic cascade and has been shown to induce the expression of SLUG as well as a number of other important neural crest genes (Aybar et al., 2003). To investigate whether neural crest cell formation is perturbed in WSTF knockdown embryos, expression of SNAIL and SLUG were analyzed by whole mount in situ hybridization in unilateral WSTF knockdown embryos (Fig. 5; representative embryos shown from a sample of 5–6 embryos for each stage and probe). Expression of both SNAIL and SLUG is unaffected on the injected side of unilaterally-injected embryos during early neurulation (stage 15; Fig. 5A and E), suggesting that neural crest induction is unaffected

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by WSTF depletion. In contrast, both SNAIL and SLUG expression patterns within the branchial arches are severely perturbed in WSTF-MO unilaterally-injected embryos later in development (Fig. 5B and C, F and G). This suggests that some neural crest cells either fail to successfully migrate to the branchial arches, or are not successfully maintained following migration. Histological sections of the branchial arches of unilateral WSTF knockdown embryos highlight the decreased expression of both SNAIL and SLUG on the WSTF knockdown side (particularly in the interior structures of the arches), and also reveal significant morphological malformations of the branchial arches on the injected side (Fig. 5D and H). These malformations include changes in shape, size and number of branchial arches in the WSTF MO-injected side, including the apparent loss or fusion of

Fig. 5 – WSTF knockdown embryos display proper neural crest induction, but fail to maintain normal neural crest in later development. A–H: Two-cell stage embryos were injected with WSTF MO unilaterally into a single blastomere and analyzed by whole mount in situ hybridization at neural groove stage (stage 15) to display expression pattern of neural crest genes SNAIL and SLUG. Both genes are expressed at the neural crest boundary. WSTF knockdown has no effect on the stage 15 expression of neural crest genes SNAIL (A) and SLUG (E) (dorsal views, anterior at top). Identically treated embryos analyzed by whole mount in situ hybridization at tailbud stage (stage 30; side views of heads, anterior at top) display perturbed expression of neural crest genes SNAIL (B and C) and SLUG (F and G) on the WSTF-knockdown side (C and G). 60-lm sections through the branchial arches of unilateral WSTF knockdown embryos (anterior at top) display diminished expression of SNAIL (D) and SLUG (H) on the injected side, particularly in the internal structures (light blue staining). Arches are labeled in B– D and F–H. Asterisks indicate the relative position of missing/fused arch number 3 on the sides derived from the WSTF MOinjected blastomeres. Dashed lines in B and F indicate the plane of sectioning of D and H, respectively. Scale bars: 1 mm (A, E), 0.5 mm (B, C, F, G) and 0.1 mm (D, H).

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arch number 3 in 5/6 sectioned embryos, and the sixth animal with no detectable arches on the injected side at all (Fig. 5C and D, G and H, and data not shown). These results strongly suggest that while neural crest induction is intact in WSTF knockdowns, the resulting neural crest cells exhibit either abnormal migration or maintenance, leading to a cascade of morphological defects.

2.5. WSTF knockdown embryos display a specific pattern of increased apoptosis The results described above indicate failures of CNS development and neural crest migration or survival in WSTF knockdowns. To determine whether the net loss of neural tissues and neural crest might be due to increased apoptosis in these tissues in our WSTF knockdown embryos, we performed a whole mount Terminal deoxynucleotidyl transferase dUTP nick end-labeling (TUNEL) assay to visualize apoptosis in situ. Anti-WSTF MO was injected at the one cell stage post-fertilization, resulting in global knockdown of WSTF within developing embryos. The TUNEL assay reveals

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an increase in apoptosis in specific regions of the WSTF knockdown embryos when compared to the controls (Fig. 6; representative embryos shown from a sample of 15 control embryos and 18 knockdown embryos at stages 31– 38). We observe increased staining at the olfactory placode and a striking increase in regions flanking the anterior dorsal midline at early tailbud stage (stage 33). This aberrant increase of apoptosis within the hindbrain is the same region where cranial neural crest cells begin their migration to the branchial arches (Smith et al., 1997), and the midhindbrain boundary is also a region that exhibits particularly robust WSTF expression beginning at stage 32 (Cus et al., 2006). A transverse section of the affected region displays punctate staining in the peripheral tissues, lateral and ventral to the hindbrain and appears to be beneath the overlying epidermal ectoderm (Fig. 6D). This punctate staining follows a pattern that is strikingly similar to the path of cranial neural crest migration and may offer an explanation for the decrease of neural crest cells present within the branchial arches of our WSTF knockdown embryos (Smith et al., 1997).

Fig. 6 – WSTF knockdown embryos display increased apoptosis of specific stages of development. A: TUNEL assay with a representative control embryo at tailbud stage showing no detectable levels of apoptosis (Stage 33; lateral view, anterior at left). B: A representative tailbud stage embryo injected with WSTF morpholino at the one-cell stage displaying a spatial increase in apoptosis in the hindbrain, olfactory placode and lateral anterior tissues. C: Side-by-side dorsal view of control embryo (left) with WSTF knockdown embryo (right) at tailbud stage, showing an increase in apoptosis in the hindbrain of the WSTF knockdown embryo compared to control (Stage 33; dorsal view, anterior at top). D: Transverse section of WSTF knockdown embryo (at a plane indicated by the dashed line in B) displays punctate staining in the peripheral tissues, lateral and ventral from the hindbrain, beneath the overlying epidermal ectoderm (Stage 33; sectional view, anterior at top). STD: standard control morpholino; MO: WSTF morpholino; nt: neural tube; nc: notochord. Scale bars: 0.5 mm (A and B), 0.1 mm (C and D).

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3.

Discussion

We have examined the specific roles of WSTF during development in order to better characterize the contribution of WSTF haplodeficiency to the clinical manifestations of WS. We show that WSTF has a crucial role in neural development and proper neural crest cell function in X. laevis. WSTF knockdowns display severe defects in neural development, and reduced expression of the neural-specific marker NCAM and reduction of PAX2 and EPHA4 within the mid and hindbrain. The expression of WSTF in neural crest cells has been well characterized (Cus et al., 2006). A number of genes have been implicated in the complex sequence of neural crest cell formation and function. In Xenopus, the neural crest-specific transcription factor SNAIL has been shown to precede SLUG and to induce its expression, as well as the expression of a number of other neural crest markers (Aybar et al., 2003). We show that knockdown of WSTF does not affect the expression of either SNAIL or SLUG during the initial stages of neural crest formation. This indicates that normal levels of WSTF are not required for neural crest induction, but does not rule out the possibility that some WSTF is essential for this process. Since some WSTF (likely maternally deposited) remains in our knockdowns, this could provide a sufficient level of WSTF for neural crest induction. However, WSTF knockdown embryos do display decreased expression of both SNAIL and SLUG within the branchial arches at early tailbud stage, suggesting WSTF is required for the normal migration and/or maintenance of the neural crest cells. Strikingly, WSTF knockdown results in significant misregulation of two signaling molecules crucial in neural development. The first of these is Bmp4, a mitogen required for the proper formation of many neural structures that are also dependent on Shh signaling. Bmp4 and Shh act in an antagonistic fashion to pattern and shape the developing neural tube, vertebrate eye and brain (Bakrania et al., 2008; Hu et al., 2004; Muller et al., 2007). BMPs, and Bmp4 in particular, inhibit ectoderm from assuming a neural tissue fate. Bmp4 not only suppresses neural tissue formation from ectoderm, but can also induce epidermal tissue determination (Hemmati-Brivanlou and Thomsen, 1995; Wilson and HemmatiBrivanlou, 1995). Shh is a developmental mitogen essential for proper neural patterning (reviewed in Patten and Placzek, 2000). Shh is believed to facilitate normal neural tissue formation by three main processes. First, Shh is known for its role in the ventral patterning of the neural tube (reviewed in Patten and Placzek, 2000). Second, Shh promotes cell differentiation by facilitating cell cycle exit (Agathocleous et al., 2007). Lastly, Shh is an essential survival factor for a number of neural cells by inhibition of the p53-dependent apoptotic pathway (Ahlgren and Bronner-Fraser, 1999; Ahlgren et al., 2002; Prykhozhij, 2010). In addition to its role in patterning of neural tissue, a classic example of Shh’s role in neural crest function is holoprosencephaly, a developmental syndrome characterized in humans by head and facial malformations, including cyclopia, resulting from haplodeficiency of SHH (reviewed in Murdoch and Copp, 2010). Reduced SHH results in decreased cell proliferation in the neural tube and neural crest, as well as smaller head size, defining a role for SHH in the survival of

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cranial neural crest (Ahlgren and Bronner-Fraser, 1999; Ahlgren et al., 2002). We show that WSTF knockdown results in a reduction of SHH expression, as determined by both in situ hybridization and realtime RT-PCR at a number of developmental stages. Additionally, a whole mount TUNEL assay reveals that WSTF knockdowns exhibit an increase in apoptosis within the hindbrain and along a path that closely mimics the initial route of cranial neural crest migration to the branchial arches (Smith et al., 1997). Shh’s role in cranial neural crest survival (Ahlgren et al., 2002) could explain our results in which both SNAIL and SLUG expression is unaffected during neural crest induction, but is then severely decreased later in development in our WSTF knockdowns, consistent with a failure in maintenance/survival of neural crest. The increased and aberrant expression of BMP4 in our WSTF knockdowns, along with the absence of WSTF expression within the notochord of normally developing embryos, suggests that the effect on SHH expression is likely to be downstream of the altered BMP4 expression. In two classic experiments aimed to investigate the function of BMP4, BMP4 was overexpressed in the animal hemisphere of fertilized Xenopus embryos (Dale et al., 1992; Jones et al., 1996). In these studies WSTF knockdowns display reduced expression of specific neural markers, including loss of PAX2 in the MHB, EPHA4 in rhombomeres 3 and 5, and SOX2 throughout the brain, eye and lateral line placodes. WSTF knockdowns also show reduction of the ear1y neural development and neural ectoderm marker NCAM. These neural defects could be due to an increase in ectodermal BMP4 expression in our WSTF knockdowns and may result in secondary malformations of the developing mesoderm from which the SHH-producing notochord and somites (expressing MRF4) are derived. Whether the effect of WSTF on BMP4 or SHH is direct or indirect remains to be determined. However, previous work in our lab has shown that ISWI, WSTF’s binding partner in the WICH complex, is present at the BMP4 gene during Xenopus embryonic development (Dirscherl et al., 2005). In addition, ISWI knockdowns exhibit BMP4 and SHH expression levels similar to those in WSTF morphants (Dirscherl et al., 2005). Taken together, these results suggest a role for the WICH complex as a transcriptional repressor of BMP4. Pluripotent embryonic stem cells are characterized by an abundance of dynamic euchromatin that undoubtedly requires an enormous amount of input and coordination of chromatin remodelers for proper differentiation or maintenance of pluripotency (Fazzio and Panning, 2010; Serra et al., 2007). Neural crest cells are extremely versatile progenitor cells that must maintain their pluripotency well into embryonic development. WSTF expression has been detected within a number of neural progenitor cells during Xenopus morphogenesis (Cus et al., 2006). Our study is the first to suggest that neural crest formation is initiated normally in embryos with depleted WSTF levels, however, these neural crest cells are not able to maintain their proper function or contribute normally to the formation of the branchial arches later in development. Here we show the misregulation of a number of well-characterized developmental morphogens and tissue markers and propose a mechanism in which aberrant BMP4 expression leads to a disruption of mesoderm

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differentiation which severely affects neural crest cell migration and/or maintenance. This effect on neural crest and mesoderm formation could account for a number of the symptoms that occur in WS patients. Development of the branchial arches, which are highly dependent on proper neural crest function, gives rise to many structures that, if malformed, could explain a number of the symptoms present in WS. WSTF deletions in mice have recapitulated both the craniofacial and the heart defects seen in WS patients, further implicating WSTF haplodeficiency in the development of WS (Ashe et al., 2008; Yoshimura et al., 2009). Mice that are homozygous for a WSTF mutation that results in a single amino acid change (L733R) and reduced stability of the mouse WSTF protein exhibit altered skull shape, including a protruding forehead, shorter snout, flattened nasal bone, and upward curvature of the nasal tip (Ashe et al., 2008). This craniofacial development is strikingly similar to many of the facial features of WS patients and suggests that WSTF deficiency may contribute to the characteristic ‘‘elfin’’ features seen in individuals with WS. WSTF knockout mice (wstf–) are small, show significant heart defects and die just days after birth. 10% of heterozygous WSTF+/– neonatal mice also display specific heart defects similar to those seen in WS patients, such as altered structure, atrial and ventricular septal defects, and hypertrophy of ventricles (Yoshimura et al., 2009). This result strongly correlates with the malformation of the branchial arches in our morphant embryos, particularly branchial arches three and four, from which a number of cardiac structures affected in WS are derived (Hutson and Kirby, 2007; Safa and El Ghazal, 2011). This result could also explain the infantile hypercalcaemia observed in WS patients. The branchial apparatus contributes to development of the thyroid and the ultimobranchial bodies, both of which produce calcitonin to regulate blood calcium levels (Varga et al., 2008). Defects in these structures could contribute to the increased blood calcium levels seen in WS individuals, this is also consistent with the increased incidence of hypoplasia of the thyroid gland in WS patients (Manley and Capecchi, 1998; Selicorni et al., 2006). Neural crest cells also contribute to the generation of tissues of the eyes including the cornea endothelium, stroma, and muscles of the iris (Grocott et al., 2011; Iwao et al., 2008). Many WS individuals do exhibit ocular defects, including small eye openings, a stellate iris pattern, puffiness around the eyes, higher occurrence of blue pigmentation, strabismus or esotropia (cross-eyes), and abnormal extraocular muscle anatomy (Ali and Shun-Shin, 2009; Winter et al., 1996). Although it is almost certainly independent of neural crest, we also show that the pronephric kidney is reduced and poorly developed in our knockdowns, with accompanying reduced expression of PAX2, which adds to the number of effected mesoderm derived structures in these knockdowns. Interestingly, WS patients display a significantly high incidence of renal and urinary abnormalities including the presences of cysts, narrowing of the renal arteries and hypoplasia (Biesecker et al., 1987; Pober et al., 1993; Sugayama et al., 2004). We note that WSTF knockdowns in Xenopus result in phenotypes that are more severe than those observed in human WS patients or in heterozygous WSTF+/wstf- mice (homozygous deletions in mice are lethal shortly after birth),

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suggesting that redundant mechanisms may partially alleviate the effects of WSTF haploinsufficiency in mammals (Ashe et al., 2008; Yoshimura et al., 2009). Intriguingly, two other genes contained in the WS deleted region, GTF2I and GTF2IRD1, are also expressed in neural crest-derived tissues and heterozygous deletions of either gene in mice results in craniofacial defects (Enkhmandakh et al., 2009). This suggests other genes in the region may also play a role in neural crest function and have additive effects on the complexity of the WS phenotype. Additional studies will be needed in order to better comprehend the role of WSTF during vertebrate development. This study furthers our understanding of the role of ATP-dependent chromatin remodeling complexes during vertebrate development as well as offer a better understanding of a number of WS symptoms that may be attributable to haplodeficiency of the WSTF gene.

4.

Materials and methods

4.1.

Generation of Xenopus embryos

Female Xenopus were induced to ovulate by injection of 600 ll of human chorionic gonadotropin (1000 USP units/ml, Intervet, Millsboro, DE) into the dorsal lymph sac. Testes were obtained by surgical excision from a male frog euthanized in 0.06% benzocaine for 45 min. Eggs were squeezed from induced females 12–18 h after induction of ovulation. Freshly laid eggs were immediately exposed to dispersed Xenopus testis in 0.1X Marc’s modified Ringer’s (MMR) (standard protocols as per Sive et al., 2007). Fertilized embryos were then stripped of jelly coat by exposure to 2% L-cysteine for 3–5 min. All husbandry and handling of adult X. laevis was performed in compliance with our approved IACUC #181339–4.

4.2.

Microinjection of morpholino oligonucleotides

Single-stranded oligonucleotides (morpholinos) were generated by Gene Tools (Philomath, Oregon). These morpholinos are complementary to the 5 0 untranslated region of WSTF messenger RNA upstream of the translation start site, and are labeled with 3 0 -carboxyfluorescein dye. As a negative control for injection, an inverse sequence or a standard control morpholino was used. The corresponding morpholino sequences are: WSTF-MO: 5 0 GCTTCTCGTGGGATGATAGTCCC GC3 0 , inverse control MO (INV-MO): 5 0 CGCCCTGATAGTAGGGT GCTCTTCG3 0 , standard control MO (STD-MO): 5 0 CCTCTTACC TCAGTTACAATTTATA3 0 (all MOs have a 3 0 carboxyfluorescein label). Xenopus embryos were obtained and dejellied as described in Section 4.1 and transferred to mesh bottom Petri dishes filled with 6% Ficoll in 0.1X MMR solution. 5 nL of a 1 mM morpholino solution was injected into either 1 or 2 cell stage dividing embryos (42 ng of morpholino per cell, resulting in either global, or unilateral WSTF morphant embryos) with glass needles made using a Flaming/Brown micropipette puller (Sutter Instruments, Novato, CA) and a KITE-L micromanipulator (World Precision Instruments, Sarasota, Florida). Morpholino was delivered to embryos by applying 30 ms bursts of 8 psi to needles using a MPPI-2 pressure injector (Applied Scientific Instrumentation, Eugene, OR). After

MECHANISMS OF DEVELOPMENT

injections, embryos were transferred to 0.1X MMR. At 17 or 32 h post-fertilization, gross morphology of the embryos was observed and photographed. Within 24 h post fertilization, injected embryos were identified by fluorescence illumination of injected morpholinos and uninjected embryos were discarded. Embryos were collected at specified times and used for either total RNA or protein extractions or were fixed in MEMFA (0.1 M Mops pH 7.5, 2 mM EGTA, 1 mM MgSO4, 3.7% formaldehyde) for in situ hybridization or sectioning.

4.3.

In situ hybridization and TUNEL assay

DNA templates were linearized with restriction enzymes, and digoxigenin-labeled sense and antisense RNA probes were transcribed in vitro from templates using Ambion Megascript in vitro transcription kits (Megascript T7TM, Megascript Sp6TM, Applied Biosystems/Ambion, Austin, Texas). The protocol was altered in the following ways: all Megascript dNTPs were diluted to 10 lM concentrations and half of the suggested RNA polymerase concentration was used. Xenopus embryos were generated as described in Section 4.1. Embryos were collected at indicated stages and fixed in MEMFA according to Sive et al. (2007). Whole mount embryos were then hybridized with in situ hybridization probes according to Sive et al. (2007); however, for some probes the RNaseA step was omitted. BM purple (Roche, Mannheim Germany) or NBT/BCIP (Promega, Madison, Wisconsin) were used as the substrate for the alkaline phosphatase conjugated to an anti-digoxigenin antibody (Roche, Mannheim Germany). For the TUNEL assays, albino embryos were collected at indicated stages and fixed in MEMFA for two hours. A whole mount TUNEL procedure from the Harland Lab was followed (detailed in Hensey and Gautier, 1998). Briefly, embryos were bleached for 1 h under light and washed in PBS. Terminal ends were labeled with Digoxygenin-dUTP (Roche, Mannheim Germany) using Terminal Deoxynucleotidyl Transferase (TdT) (Invitrogen, Carlsbad, California) overnight at room temperature. The reaction was stopped with 65 °C washes in PBS containing 1 mM EDTA. Anti-digoxygenin AP antibody incubation was done in MAB (100 mM Maleic acid, 150 mM NaCl) containing 2% BMB blocking reagent (Roche, Mannheim Germany) overnight at 4 °C. Following extensive washes in MAB, the color reaction was performed using NBT/BCIP (Harland, 1991).

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slides (Mayer’s fixative) (Humason, 1972). Sections were stained using Ehrlich’s hematoxylin and counterstained in Eosin following the protocols of Humason (1972). Coverslips were applied using Permount (Fisher Scientific, Pittsburgh, PA). Color images were captured using a Spot digital camera (Diagnostic Instruments, Inc., Sterling Heights, MI).

4.5.

Total RNA extraction and RT-qPCR analysis

Total RNA samples were isolated from injected embryos at various times post fertilization. Ten whole embryos were homogenized in 200 ll Trizol reagent (Invitrogen, Carlsbad, California) using a mortar and pestle and the RNA was isolated following manufacturer’s recommendations. RNA pellets were re-suspended in 100 ll RNase-free water. Turbo DNase (Ambion, Austin, Texas) was used to remove any genomic DNA contamination in the RNA preparations. cDNA generation and qPCR were performed using the one-step SYBRÒ Green Quantitative RT-PCR Kit (Sigma–Aldrich, St. Louis, Missouri) in a Smart Cycler (Cepheid, Sunnyvale, California) system.

4.6.

Total protein extraction and Western blot analysis

10–20 embryos were collected and homogenized via mortar and pestle in Laemmli sample buffer. Homogenate was heated at 95 °C for 5 min, centrifuged at 13,000 rpm for 1 min and the supernatant was stored at 80 °C. Total protein extracts were separated by 10% SDS polyacrylamide gel electrophoresis, blotted onto a nitrocellulose membrane (Whatman, Dassel, Germany) and blocked in Odyssey blocking buffer (Licor, Lincoln, Nebraska). The membrane was probed with the primary antibody in Odyssey blocking buffer at room temperature overnight. Primary antibody against Xenopus WSTF was generously provided by Dr. Paul Wade (NIEHS). A b-actin monoclonal antibody (Abcam ab8224, Cambridge, Massachusetts) was used for a loading control. IR-dye conjugated (800CW) goat anti-mouse secondary antibodies (Licor, Lincoln, Nebraska) were used for secondary antibodies. The Odyssey infrared imager (Licor, Lincoln, Nebraska) was used to visualize the probed membranes.

Acknowledgements 4.4.

Histology

Embryos were fixed in MEMFA and stored at 20 °C in 100% methanol. Embryos were gradually rehydrated by incubating in 1X PBS containing 75%, 50%, 25% methanol and in 1X PBS for 5 min each. They were then embedded in 15% gelatin (Bloom 300, Sigma–Aldrich, St. Louis, Missouri) in PBS (135 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.4 mM KH2PO4, pH 7.2) and fixed in 10% neutral buffered formalin (3.7% formaldehyde) (BDH, West Chester, PA) for 36–48 h at 4 °C. The gelatin blocks were rinsed in PBS and 60 lm sections were cut using a Leica VT1000P vibratome (Richmond, Illinois). Other fixed embryos were dehydrated though a graded series of ethanol and xylene washes, embedded in Paraplast Plus (Oxford Labware, St Louis, MO) and serially sectioned at a thickness of 7 lm. Sections were collected on albumin-subbed

Template DNAs for SLUG and SNAIL were generously provided by Peter Walentek in Dr. Martin Blum’s laboratory (University Hohenheim, Stuttgart, Germany). BMP4 and SHH DNA templates were generously provided by Dr. Richard Harland (University of California, Berkley), and fully transcribed MRF4 probe was provided by Dr. Tim Hinterburger (University of Alaska Anchorage). The NCAM and SOX2 RNA probes were developed at the Cold Spring Harbor course on Developmental Biology of Xenopus. Primary antibody against Xenopus WSTF was generously provided by Dr. Paul Wade (NIEHS). This work was supported by NIH EY016029 and Whitehall Foundation 2007-08-79 to JEK, and NIH EY09844 to JJH. CB was also supported by NIH P20RR016466. This paper is dedicated to the memory of Dr. Marietta Dunaway (1952-2011).

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Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/ j.mod.2012.06.001.

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