Role of Chd7 in Zebrafish: A Model for CHARGE Syndrome

Abstract

CHARGE syndrome is caused by mutations in the CHD7 gene. Several organ systems including the retina, cranial nerves, inner ear and heart are affected in CHARGE syndrome. However, the mechanistic link between mutations in CHD7 and many of the organ systems dysfunction remains elusive. Here, we show that Chd7 is required for the organization of the neural retina in zebrafish. We observe an abnormal expression or a complete absence of molecular markers for the retinal ganglion cells and photoreceptors, indicating that Chd7 regulates the differentiation of retinal cells and plays an essential role in retinal cell development. In addition, zebrafish with reduced Chd7 display an abnormal organization and clustering of cranial motor neurons. We also note a pronounced reduction in the facial branchiomotor neurons and the vagal motor neurons display aberrant positioning. Further, these fish exhibit a severe loss of the facial nerves. Knock-down of Chd7 results in a curvature of the long body axis and these fish develop irregular shaped vertebrae and have a reduction in bone mineralization. Chd7 knockdown also results in a loss of proper segment polarity illustrated by flawed efnb2a and ttna expression, which is associated with later vascular segmentation defects. These critical roles for Chd7 in retinal and vertebral development were previously unrecognized and our results provide new insights into the role of Chd7 during development and in CHARGE syndrome pathogenesis.

About the Authors:

Shunmoogum A. Patten

Contributed equally to this work with: Shunmoogum A. Patten, Nicole L. Jacobs-McDaniels

Affiliations Sainte-Justine Hospital Research Center, Montreal, Quebec, Canada, Faculty of Dentistry, University of Montreal, Montreal, Quebec, Canada

Nicole L. Jacobs-McDaniels

Contributed equally to this work with: Shunmoogum A. Patten, Nicole L. Jacobs-McDaniels

Affiliation: Department of Biology, Syracuse University, Syracuse, New York, USA

Charlotte Zaouter

Affiliation: Sainte-Justine Hospital Research Center, Montreal, Quebec, Canada

Pierre Drapeau

Affiliation: Department of Pathology and Cell Biology, Faculty of Medicine, University of Montreal, Montreal, Quebec, Canada

R. Craig Albertson

Affiliations Department of Biology, Syracuse University, Syracuse, New York, USA, Department of Biology, University of Massachusetts, Amherst, Massachusetts, USA

Florina Moldovan

* E-mail: [email protected]

Affiliations Sainte-Justine Hospital Research Center, Montreal, Quebec, Canada, Faculty of Dentistry, University of Montreal, Montreal, Quebec, Canada

Introduction

Mutations in the CHD7 gene (NM_017780) in 8q12.1 were identified as causative for CHARGE (Coloboma, Heart defects, Atresia of the choanae, Retarded growth and development, Genital hypoplasia, Ear anomalies; OMIM # 214800)) syndrome [1], [2], [3], [4], [5]. Heterozygosity for nonsense, deletion or missense CHD7 mutations is estimated to occur in 60–80% of patients with CHARGE syndrome; these mutations are distributed throughout the coding sequence and do not appear to be correlated with specific aspects of the clinical phenotype [6], [7], [8]. The majority of CHD7 mutations identified thus far are de novo; however, evidence for germline mosaicism has been suggested for families with multiple affected siblings [9] . Children affected by CHARGE syndrome have a variable association of features including ocular deficits, heart malfunctioning, olfactory dysfunction, retarded growth, vestibular dysfunction, cranial nerve anomalies and intellectual disability [2], [4], [10]. All of these characteristics have variable degrees of penetrance, with some being present in virtually all CHARGE patients, whereas others are less frequently observed [11]. Although there is clear evidence that mutations in CHD7 gene are causative in CHARGE syndrome, the pathogenic mechanisms elicited by these mutations that lead to organs and systems dysfunction are not fully understood.

Chd7 is one of nine members of the chromodomain helicase DNA- binding (Chd) domain family of ATP-dependent chromatin remodeling enzymes [12]. It consists of functional domains such as a chromatin organization modifier, SNF2-related helicase/ATPase and BRK [12], [13]. Recently, DNA-binding sites on chromatin have shown that chd7 binding is correlated to areas of mono- and dimethylated lysine 4 of histone H3 [14], [15]. The Chd7 Drosophila ortholog, Kismet down-regulates transcriptional elongation by RNA polymerase II through the recruitment of ASH1 and TRX and may be involved in the maintenance of stem cell pluripotency by regulating methylation of histone H3 lysine 27. Chd7 is also implicated in cell fate specification of mesenchymal stem cells [16] . During osteoblast and adipocyte differentiation, Chd7 forms a complex with NLK, SETDB1 and PPAR-γ, then binds to histone H3 at PPAR-γ target promoters and suppresses ligand-induced transactivation of PPAR-γ target genes, which leads to a change in cell fate [16] . Recently, Chd7 has been shown act synergistically with PBAF to promote neural crest gene expression and cell migration [17]. Thus, Chd7 is thought to play a variety of essential roles during development in many species.

The understanding of cause and pathogenesis of many complex human diseases has been improved through the study of model organisms. Zebrafish are a well-established model used to study developmental biology because of their accessibility, optical transparency and rapid development. Over the past decade, the utility of this model organism in investigations of human health and disease has become more explicit. Major advantages of this vertebrate model include a high degree of homology to human genes, as well as conservation of developmental pathways. Zebrafish chd7 gene is located on chromosome 2 and alignment of the zebrafish genomic sequence with mouse Chd7 and human CHD7 shows the gene structure is conserved across species [18]. Given that the zebrafish chd7 gene is highly similar to the human CHD7 gene, elucidating roles for Chd7 in zebrafish development could significantly contribute to our understanding of the function of Chd7 in CHARGE syndrome pathogenesis. Therefore, we sought to utilize zebrafish to assess the pathogenic effect of the loss of function of Chd7 and understand the role of the zebrafish homolog of the mammalian Chd7 during development. We show that chd7 is robustly expressed in developing zebrafish embryos, particularly in the retina, hindbrain and tail bud. Loss of function of Chd7 results in several defects including abnormal neural development, curvature of the long axis of the body, abnormal cranial neural crest (CNC) development, otolith anomalies, smaller eye and pericardial edema. These defects are similar to the major congenital anomalies associated with human CHD7 mutations in CHARGE syndrome. Finally, we reveal for the first time a critical role of Chd7 in retinal organization and bone mineralization.

Results

Chd7 transcript is broadly expressed in developing zebrafish

To begin to explore the role of Chd7 in developing zebrafish, we first analyzed the developmental expression of chd7 mRNA. In situ hybridization results showed that chd7 was expressed relatively ubiquitously throughout early zebrafish embryogenesis (Fig. 1 A–C) until around the 13 somite stage of development when more discrete expression in the eye primordium, and the perimeter of the somites was observed (Fig. 1 D). As somitogenesis continued, chd7 became highly expressed in the retina as well as in the brain, somites, and tailbud of the embryo (Fig. 1 E–F). Finally at 48 hpf, chd7 expression in the tail began to diminish, however expression in the brain and in the eye remained strong (Fig. 1 G). The sense probe did not show any in situ staining suggesting that there was no non-specific hybridization (Fig. 1 E′).

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Figure 1. Chd7 expression patterns during zebrafish embryogenesis.

Chd7 was expressed ubiquitously throughout the embryo during epiboly (A), at the 4 somite stage (B), and at the 8 somite stage (C). (D) At the 13 somite stage, expression remained relatively ubiquitous, but with stronger expression noted within the retina and at the perimeters of developed somites. (E) At the 18 somite stage, tissue-specific expression was observed in the eyes, brain, somites, and tailbud. This expression pattern remained through 24 hours post fertilization (F). (E′) No staining occurred when hybridized with a chd7 sense probe. (G) At 48 hours post fertilization, chd7 expression began to diminish from the body of the zebrafish but remained in the eye and in the brain. e, eye; ep, eye primordium; mb, midbrain; hp, head primordium; s, somite; t, tail; tb, tailbud; y, yolk; ye, yolk extension.

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Loss of function of Chd7 leads to several and widespread morphological abnormalities

Next, we depleted zebrafish embryos of Chd7 protein using an antisense morpholino oligonucleotide (MO) designed against the translation initiation sites of chd7 mRNA. Embryos injected with chd7-MO exhibited phenotypes marked by curvature of the long axis of the body, a flattening of the head, abnormal tail fins, smaller eyes and heart defects (n = 12, A total of 498 fish used; Fig. 2, 3 A–C). These defects were observed in a dosage-dependent manner (Figure S1). As embryos were injected with increasing concentrations of chd7-MO, the resulting defects became more severe (Figure S1 C–D)

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Figure 2. Chd7- MO injections and mRNA rescue experiments.

Control-MO injected (E) chd7-MO+chd7-mRNA co-injected (F) zebrafish showed no phenotypic defects at 48 hpf and were comparable to wild type zebrafish (A) at the same age. Embryos injected with 2 ng/nl chd7-MO (B–C), or 2 ng/nl (D) chd7-SBMO showed several developmental defects.

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Figure 3. Knockdown of chd7 affects eye and heart development.

(A–B) Representative images of eye phenotypes induced by injection of chd7-MO in zebrafish embryos (B) compared with control embryos (A) at 72 hpf. Lateral view with anterior to right. Chd7 morphants had a reduced ocular size. Chd7-MO-injected zebrafish exhibited cardiac anomalies such as dysmorphic heart and tube-like heart shape (D) compared with control embryos (C). Insets in (C) and (D) represent an illustration of the heart phenotype in control or chd7 morphant embryos, respectively. (E–F) Bar graphs illustrating the prevalence of eye (E) and heart (F) phenotypes in wild-type (white bar), chd7-MO (black bar), control-MO-injected embryos (grey bar) and mRNA rescue (dark grey bar). ***<0.001, *<0.01. ***<0.001, *<0.01.

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To exclude the possibility that this phenotype may have resulted from non-specific or mistargeting effects of the MO, we also designed and injected zebrafish embryos with a chd7 splice-blocking morpholino (SBMO) and control 5-bp mismatch MO. We recently showed by RT-PCR analysis that the morpholino-targeted region exhibited abnormal splicing of chd7 in embryos injected with chd7 SBMO and normal splicing in uninjected and control-MO injected embryos [19]. Embryos injected with the chd7 SBMO exhibited morphological defects similar to those observed in the chd7-MO morphants (Fig, 2 D). On the other hand, all control MO-injected larvae developed normally (n = 12, 436 fish; Fig. 2 E and Fig. 3) and no apparent phenotype was observed when compared to untreated wild-type embryos (n = 12, 510 fish; Fig. 2 A). To further confirm that the lack of Chd7 underlies all the phenotypic alterations observed in chd7-MO zebrafish, a rescue experiment was performed. We found that expression of chd7 mRNA which had no complementary sequence to chd7-MO can correct all the developmental defects (n = 6, 249 fish; Fig. 2 E) and no apparent phenotype was observed when compared to control-MO fish (Fig. 2). Thus, the amount of newly-synthesized Chd7 coded by the rescuing mRNA in the initial stages was sufficient to prevent the developmental anomalies imparted by the chd7-MO. Altogether these findings were consistent with a reduction in Chd7 expression and they suggest that the phenotype observed with chd7-MO-injected embryos was a specific and due to a direct result of Chd7 protein depletion.

We focused the analysis of the loss of function of Chd7 in embryos injected with 2 ng/nl of chd7-MO. At 48 hpf, we found that chd7-MO-injected (2 ng/nl) embryos exhibited a small eye phenotype (Fig. 3B,E) compared to control-MO (Fig. 3A,E) and wild-type fish (Fig. 3E). We observed that the hearts of chd7-MO-injected embryos were developmentally impaired and have signs of severe pericardial edema compared to controls and wild-type fish (Fig. 3 F). The atrial and ventricular chambers in these fish can be distinguished (Fig 3. D), but these compartments were dysmorphic and more tube-like (inset, Fig. 3 D) than the well-defined and tightly looped chambers of control-MO injected fish and wild-type fish (Fig. 3C). Injection of chd7 mRNA significantly rescued the morpholino eye and heart phenotypes (Fig. 3E–F).

At 48 hpf, chd7-MO-injected (2 ng/nl) embryos also displayed a very specific defect in otolith formation (n = 6, 214 fish; Fig. 4B–D). We observed that 63% of the morphants exhibited otoliths that were asymmetric in size (135/214; p = 0.006, ANOVA; Fig. 4 B,D), and 15% possessed only one otolith (32/214; p = 0.008, ANOVA; Fig. 4 C–D). The enlarged or single remaining otolith was often shaped irregularly as opposed to the round and smooth otoliths in control-MO (n = 6, 249 fish; Fig. 4 C), wild-type embryos or mRNA rescue embryos (Fig. 4-D). In addition, the general size and morphology of the ear and the formation of semicircular canals were affected in morphants with a single otolith. Fish with defective otoliths also had a pronounced curvature of the body axis, usually remained on their sides at the bottom of their dish, and exhibited a circling swimming behavior upon a touch (tail tap) or acoustic (dish tap) stimulation (data not shown).

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Figure 4. Knockdown of chd7 affects otolith development.

(A) The otic vesicle of control-MO-injected fish contains two otoliths, by contrast, larva treated with a morpholino against chd7 (B,C) had smaller otoliths (*) or only a single otolith (Arrow). Lateral views with anterior to the left. Ot, otoliths (D) Bar graphs illustrating the prevalence of otolith phenotypes in wild-type (white bar), chd7-MO (black bar), control-MO-injected embryos (grey bar) and mRNA rescue (dark grey bar). ***<0.001, *<0.01.

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Chd7 plays critical roles in primary axis development and vertebral mineralization

Loss of function of Chd7 led to a pronounced curvature of the body axis in 25% of the morphants (n = 12; p<0.001, ANOVA; Fig. 5 A–C) compared to control-MO (n = 12; Fig. 5 A,C) and wild-type embryos (n = 12; Fig. 5 C). This phenotype was completed rescued by co-injection of chd7 mRNA (n = 6; Fig. 5 C). Recently, CHD7 gene polymorphisms were associated with susceptibility to idiopathic scoliosis in human populations [20]. We therefore sought to examine vertebral phenotypes at various developmental stages (Fig. 5 D–F). Chemical staining of the vertebrae at later stages revealed that the spine of zebrafish injected with 2 ng/nl of chd7-MO had several skeletal anomalies but exhibited no scoliotic curves (n = 6, 242 fish; Fig. 5 E,F). Notably, chd7-MO-injected (2 ng/nl) fish had smaller, irregular vertebral segments (n = 6; Fig. 5 F), wider intervertebral disc space (n = 6; Fig. 5 F) and smaller neural and hemal spines (Arrows; n = 6; Fig. 5 F). The pedicles of the neural spines were in some cases missing or fused. Mineralization can be assessed by the intensity and localization of the alizarin red S stain [21] .We observed a marked decrease in the alizarin red S staining of the vertebral segments of chd7-MO-injected fish (n = 6, 242 fish; Fig. 5 D–F) compared to control-MO ,suggesting a reduction in vertebral mineralization in chd7 morphants . These data suggest chd7 could play a critical role in vertebral development.

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Figure 5. Chd7 deficiency affects vertebral mineralization.

Lateral view of control-MO (A) and chd7-MO-injected (B) fish at 3 dpf. Chd7 morphants exhibited a slight curvature of the long body axis (B). A significant percentage of the chd7-MO-injected (2 ng/nl) fish had deformed body axis shape (C). Lateral views of chd7 morphants (lower panel) and control siblings (upper panel) at 8 dpf (D) and 14 dpf (E) showed a reduced bone mineralization. Vertebrae with reduced or no hemal or neural spines were also observed in the chd7-MO-injected fish (F; black arrow). h, hemal spine; n, neural spine. ***<0.001.

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Chd7 is required for proper somite polarity and segmental vascularization

Morpholino effects on protein abundance typically last for <7 days [22] and surprisingly, we observed post-embryonic vertebral defects in chd7-MO zebrafish. However, the skeletal malformation observed in chd7-MO injected fish (in Fig. 5) resembles the phenotype observed when somite segmentation is disrupted [23], [24]. To determine whether the abnormality in segmentation of the vertebral axis was due to developmental somite malformation, we analyzed the somite segmentation phenotype. To do so, we examined expression of the segment-polarity gene, ephrin B2a (efnb2a). In situ hybridization expression data for efnb2a revealed defects in the delineation of somite borders. To verify this defect in somite borders, a segment boundary marker, titin a (ttna), was used. Specifically, zebrafish chd7 morphant embryos showed a loss of distinct borders between individual somites at the 13 somite stage, as exhibited by efnb2a expression (146 fish;p = 0.008, t-test; Fig. 6 A–C) and ttna expression (112 fish; p = 0.006, t-test; Fig. 6 D–F). Stage matched control MO injected embryos exhibited distinct expression of efnb2a within each somite (149 fish, Fig. 6 A–B), and ttna expression marked clear boundaries between adjacent somites (143 fish, Fig. 6 D–E). Sectioning of efnb2a stained embryos revealed that this defect occurs throughout the somitic tissue in chd7 morphants (inset, Fig. 6 A–B). Therefore, chd7 appears to be important for proper segment boundary formation. These results also suggest that the skeletal malformation observed later on during development in chd7 morphants was likely caused by defective somite segmentation.

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Figure 6. Chd7 knockdown leads to somite boundary and segmental vasculature defects.

Efnb2a expression in control morphant (A) and chd7 morphant (B) zebrafish embryos at the 13-somite stage of development. Top is anterior. Insets in (A) and (B) represent coronal sections through control or chd7 morphant embryos, respectively, at the 13 somite stage. Posterior views with dorsal towards the top. Expression of ttna, a segment boundary marker reiterates the defects in segment boundary formation in chd7 morphants (E) compared to controls (D). Dorsal views with anterior to the top. Segmental vasculature patterning is defective (arrowheads) in chd7 morphants (H) compared to control morphants (G) as visualized using fli1: GFP transgenic zebrafish at 48 hpf. Sagittal views with anterior to the right. Graphs show percentages of animals exhibiting efnb2a expression defects (C), ttna defects (F), or segmental vasculature patterning defects (I). The dark gray bar represents chd7 morphants and the light gray bar represents control morphants. Scale bar: 200 µm. ***<0.001, **<0.01.

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Furthermore, since efnb2a is involved in arterial-venous differentiation [25] and the development of intersegmental vasculature is dependent on proper segmentation of the presomitic mesoderm (PSM), we utilized a fli1: GFP transgenic zebrafish line to test for defects in vascular development caused by Chd7 knockdown. With this resource, we observed aberrant vascular organization. After 48 hours, chd7 MO injected embryos displayed improper patterning of the intersegmental vasculature along the long axis of the body (56 fish; p = 0.001,t-test; Fig. 6 G–I). Since these vessels form at the boundaries between somites [26], this phenotype is likely due to a lack of intersomite specification in chd7 morphants, and is similar to defects observed in other mutants defined by aberrant PSM segmentation [25].

Chd7 function is required for retinal organization

We next thought to further analyze the eye morphology of the chd7 morphants. To permit a detailed assessment of the chd7-MO small eye phenotype, histological analysis was performed on embryos at 72 hpf (Fig. 7). Histology revealed that retinal organization was severely disrupted in chd7-MO–injected embryos (120 fish, Fig. 7 B). Specifically, the retinas lacked the characteristic laminated structure of both control-MO (142 fish, Fig. 7 A) and wild-type embryos at 72 hpf. Immunostaining further confirmed disruption of retina formation. Retinal ganglion cells appear to be reduced and disorganized in the chd7-MO–injected embryos based on staining for the retinal ganglion marker Zn-8 (n = 5; Fig. 7 C, D). Staining with 3A10, a neurofilament marker was abnormal in that the normally recognizable stripe of expression of the photoreceptor layer was not present in the chd7-MO–injected embryos (n = 8; Fig. 7 E, F). Similarly, cone photoreceptor staining with Zpr-1 antibody was absent in chd7 morphants (Figure S2). These findings suggest a novel role of Chd7 in retinal organization and the development of photoreceptors.

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Figure 7. Chd7 plays an essential role in retinal development.

Retinal organization of control-MO embryos (A) and chd7 morphants (B) was revealed by toludiene blue staining. Compared with the highly organized cells and laminated retinal structure in control-MO fish, chd7-MO retinal cells are disorganized (A–B: Left panels). Retinal lamination defect, including rosette formation,is clearly visible in the chd7 morphants (examples of rosettes are indicated by dotted lines and arrow heads). GL, ganglion cell layer; INL, inner nuclear layer; L, lens; ONL, outer nuclear layer; R, retina; RPE, retinal pigment epithelium. Scale bar: 50 µm. Zn-8 immunoreactivity was performed to label (brown) retinal gangion cells in control-MO (C) and chd7-MO embryos (D). Scale bar: 30 µm. The expression of retinal ganglion cell-specific marker zn-8 is greatly reduced in chd7 morphants. The photoreceptor layer of control-MO (E) and chd7-MO-injected (F) embryos were stained with 3A10. Chd7 morphants lacked the photoreceptor layer. Scale bar: 5 µm.

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Chd7 is essential for correct positioning of cranial motor neurons and proper projection of facial and trigeminal motor axons

Previous studies have demonstrated a role for Chd7 in neuronal development [27], [28], [29] and patients with CHARGE syndrome often display several neurological disorders. We found that chd7 is expressed in the developing zebrafish hindbrain and loss of function of chd7 leads to flattening of the head. To determine if these molecules are required for proper neural development in this area, we examined the cranial motor neurons of chd7-MO-injected embryos using Isl1-GFP transgenic zebrafish embryos. These embryos express GFP under the control of a motor neuron-specific Isl1 promoter. Interestingly, we observed a disrupted organization of the cranial motor neurons in the chd7-MO-injected embryos (n = 9; Fig. 8 B) when compared to control-MO-injected embryos (n = 9; Fig. 8 A). Chd7-MO-injected embryos demonstrated a severe disorganization and clustering of both Va and Vp clusters of the trigeminal (nV) neurons (Fig. 8 B). In addition, loss of Chd7 resulted in a reduction of facial (nVII) branchiomotor neuron populations (n = 9; Fig. 8 B). Furthermore, the vagal (nX) motor neurons in chd7-MO-injected embryos exhibited aberrant medio-lateral positioning and cell-to-cell spacing (n = 9; Fig. 8 B). Chd7-MO-injected embryos also exhibited extremely low levels of staining of the peripherally extending axons from nV neurons when compared to control-MO-injected embryos (n = 9; Fig. 8 C, D). Furthermore, chd7-MO injected embryos exhibited severe loss of the facial sensory ganglion and the axonal extensions from nVII neurons (n = 9; Fig. 8 C, D).

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Figure 8. Chd7 is important for cranial ganglia development and proper projection of facial nerves.

Dorsal view (rostal towards the top) of confocal fluorescent composite images of hindbrain branchiomotor neurons in control-MO injected (A) and chd7-MO-injected (B) 48 hpf Isl1-GFP transgenic embryos. (C, D) Confocal fluorescent composite images showing anti-3A10 antibody-stained axons of 48 hpf embryos. The broken lines indicate the area of significant loss of VII nerves in chd7 morphants. nV, trigeminal motoneurons; Va, anterior trigeminal motoneurons; Vp, posterior trigeminal motoneurons; nVII, facial motoneurons; nX, Vagal motoneurons; V, trigeminal nerve; VII, facial nerve. Scale bar: 50 µm.

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Chd7 plays a key role in cranial neural crest development

CHARGE syndrome is thought to result from the abnormal development of the neural crest. Recently, CHD7 and PBAF have been shown to cooperate during embryonic development to promote proper neural crest specification and cell migration [17]. We sought to examine the relationship between Chd7 and CNC development in zebrafish using a Fli1-GFP transgenic zebrafish line. We injected Fli1-GFP embryos with chd7-MO at the single cell stage and then assayed for CNC defects between 34–36 hpf by counting the number of CNC segments, which should approximate pharyngeal arch number, within each embryo. We observed CNC defects in over 90% of chd7-MO-injected embryos (63 fish, Fig. 9 D). About one third of these had a reduction in the number of CNC segments, whereas the remaining animals showed more severe phenotypes characterized by the gross disorganization or absence of obvious CNC populations (Fig. 9 B, C). Disorganized CNC segments tended to possess fewer cells and lacked sharp borders between adjacent segments (Fig. 9 C). The phenotype of control-MO-injected zebrafish embryos mimicked that of wild type animals including well-defined segments and appropriate segment numbers (71 fish, Fig. 9 A). These data lend support for the function of Chd7 in proper CNC development and patterning. Whether the defects reported here are due to aberrant CNC migration or improper pharyngeal arch patterning remains to be investigated.

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Figure 9. Knockdown of chd7 results in defects in CNC development.

Fli1-GFP transgenic embryos were used for MO injections to determine whether Chd7 depletion had an effect on CNC development. Embryos were injected at the 1 cell stage and examined for effects at 34–36 hpf. (A) Control-MO injected zebrafish embryos did not exhibit defects in CNC segment shape or number. Alternatively, the majority of chd7-MO-injected embryos exhibited aberrant CNC development including an improper number of CNC segments (B), as well as severely disorganized (C) or missing CNC populations. (D) Based on the t-test comparing means, there was a significant difference in CNC defects between chd7-MO and control-MO-injected zebrafish embryos. More than half of the chd7-MO-injected fish had deformed or missing CNC segments and about one third developed an improper number of segments. ** = 0.01; ***<0.001; e, eye; mb, midbrain; pa1–5, pharyngeal arches 1–5.

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Discussion

The present results show a novel and essential role of Chd7 in retinal organization, axial patterning and bone development and mineralization. Furthermore, knockdown of zebrafish chd7 results in abnormalities similar to those described in CHARGE Syndrome, as we observed inner ear, heart, eye, cranial ganglia, neural crest and skeletal defects in chd7 morphants. Collectively, our findings demonstrate that Chd7 function is essential for proper neural and cranial neural crest development. In addition, to our knowledge, this is the first study to report the requirement of chd7 in retinal and vertebral development.

Chd7 has been implicated in spinal deformities [20] and scoliosis is one of the minor clinical features in some CHARGE patients [11]. Here, loss of function of Chd7 resulted in the curvature of the body axis, irregular vertebral segments and smaller neural and hemal spines. The chd7 morphants did not exhibit any skeletal defects reminiscent of a scoliotic adult phenotype, however, suggesting that either scoliosis in CHARGE patients is secondary to the effects of attenuated Chd7 levels, or due to primary effects that occur later in development. Our findings also showed that zebrafish embryos deficient in Chd7 mineralize vertebrae more slowly than control siblings, indicating a critical role for Chd7 in promoting mineralization during normal development. Delays in vertebral mineralization may result from a general retardation in growth and development; however, we did not detect any distinguishable differences in length between the chd7 morphants and controls at all developmental stages examined (Table S1). Interestingly, a similar delay of mineralization in zebrafish has been reported after knockdown of collagen XXVII [30]. Furthermore, in the absence of BMP signaling, zebrafish bone mineralization was delayed [31]. Chd7 has been shown to regulate BMP signaling [27], [32]. Thus, it is possible that disruption of Chd7 affected the transcription of genes that are crucial for bone mineralization. We also show that a loss of somite border identity during somitogenesis, as exhibited by efnb2a and ttna expression, is linked to a loss of organized segmental vasculature following chd7 MO injection and skeletal defects. We hypothesize that early defects in efnb2a expression lead to improper segmental patterning later in development, affecting structures such as segmental vasculature and bone mineralization. The link between the process of somitogenesis and the defects in border specification will require further investigation. For example, since efnb2a expression persists after somite epithelialization, the irregular efnb2a expression observed here could be the result of an accumulation of upstream defects occurring early in the somitogenesis signaling pathway (i.e., indirect effects). Alternatively, it is also possible that Chd7 modulates efnb2a expression in a manner that is independent of the process of somite formation (i.e., direct effects).

CHARGE syndrome patients often have microphthalmia [33], [34], [35] and here we found that loss of function of Chd7 in zebrafish resulted in a small eye phenotype. The retinas in zebrafish develop rapidly and begin differentiation by 36 hpf [36], [37]. The differentiating retinal neurons undergo terminal differentiation by 52 hpf and become functional at around 3 dpf [37], [38], [39]. A functional retina consists of a laminated neuronal tissue with the ganglion cell layer on the inside, inner nuclear layer in the middle, and the outer nuclear layer on the outside. In between these layers are the synaptic layers: the inner and outer plexiform layer [36], [40]. Notably, the retinas in chd7 morphants fail to laminate, which suggests a novel function for Chd7 in retinal patterning. Furthermore, our findings are similar to disruptions in retinal lamination as observed in the young zebrafish mutant that carries a mutation in brg1, a gene that encodes for a subunit of the Swi/Snf of ATP-dependent chromatin remodeling complexes [37]. Moreover, retinal ganglion cells were reduced and the photoreceptor layer were absent in the chd7-MO-injected embryos. These data suggest that Chd7 is required for the development of retinal cells. In Drosophila, the chd7 ortholog kismet is essential for transcription of the pro-neural factor atonal (Atoh1) and regulation of retinal photoreceptor cell development [41]. Drosophila ato mutants produce no photoreceptors in the eye [42]. Chd7 has also been recently shown to control the expression of sox9 [17]. Zebrafish sox9 mutants reveal that Sox9 is required for retinal differentiation and it also helps with retinal organization and regulates the number of photoreceptor cells [43]. Thus, it is likely that that loss of function of Chd7 disrupts the gene network that is crucial for retinal cellular development and organization.

Several ganglionic defects have been reported in CHARGE patients, including defects in the facial and vestibulo-acoustic ganglia [44], [45]. Chd7 knockdown resulted in important phenotypes relevant to cranial ganglia development, branchiomotor development, and vagal motor neuron positioning. Anomaly in facial nerves has also been demonstrated in CHARGE syndrome [5]. Loss of function of Chd7 resulted in a severe loss of facial nerves in the morphants. Altogether, these results suggest that Chd7 and downstream targets are important for neuronal development and proper axonal projections.

Neural crest cells are multipotent cell populations with the ability to migrate, leading to the formation of several key developmental structures including bones, cartilages, nerves, and connective tissues [46]. CNC, specifically, migrate into the pharyngeal arches where they play a role in the formation of facial bone, muscle, and cartilage [47]. In addition, CNC migrate into the optic vesicle and otic placode, where they play key roles in the development of muscular and skeletal elements in the eye and the inner ear, respectively [48], [49]. Some key developmental defects in CHARGE patients can be traced to aberrant CNC development, including coloboma of the eye, external ear malformations, inner ear defects and a spectrum of facial defects [6], [50], [51]. Our chd7 morphant zebrafish embryos displayed obvious defects in CNC migration, based on observations of pharyngeal arch development and number. We did not fully characterize CNC activity in the optic nerve or otic placode of our chd7 morphants, however, we did observe and quantify significant defects in eye and otic vesicle development. CHD7 is essential for the formation of multipotent migratory neural crest in humans and Xenopus [17] and recently, CHD7 was found to play a role in proper neurogenesis of the inner ear in mice [27]. Our results support a role for Chd7 in proper CNC migration and differentiation, and substantiate the use of chd7 morphant zebrafish as an in vivo animal model for CHARGE Syndrome.

The chd7 –MO-injected embryos displayed a very specific defect in otolith and semicircular canal formation and they also exhibited a circling swimming behavior consistent with vestibular dysfunction. Such inner ear malformations are very similar to those reported in CHARGE syndrome patients [52], [53], [54], [55]. In addition, CHARGE patients have been reported to have vestibular problems and hyperactivity [56], and this may be due to inner ear defects. CHD7 is necessary for proliferation of inner ear neuroblasts and inner ear morphogenesis by the maintenance of Fgf10, Otx2 and Ngn1 expression [27]. Fgf signaling and Otx genes have also been shown to be important for inner ear development in zebrafish [57], [58], [59], [60]. Thus, our zebrafish model for CHARGE syndrome can be useful to further explore the mechanisms underlying inner ear anomalies in CHARGE pathogenesis. For instance, it will be interesting to investigate whether loss of function of chd7 in zebrafish results in a downregulation of Fgf10 and Otx2 and whether overexpression of one or more of these genes could rescue chd7 –MO phenotypes.

Heart defects have been reported to be associated with CHARGE syndrome [2], [3], [5], [61], [62]. Here we report that the hearts of chd7-MO-injected embryos are developmentally impaired and have signs of severe pericardial edema. Heart anomalies described in CHARGE syndrome patients include ventricular and atrial septal defects and conotruncal defects [61], [63]. In this study, we did not fully characterize the observed heart defects in the chd7 morphants. The atrial and ventricular chambers in these fish can be distinguished, but these compartments were dysmorphic and more tube-like than the well-defined and tightly looped chambers of wild-type fish. Future studies using chd7 zebrafish morphants could help to identify components in the Chd7 regulatory network that are essential for heart development and further our understanding of one of the major anomalies reported in CHARGE patients.

In conclusion, we have examined the role of zebrafish Chd7. We provide evidence that zebrafish highly express chd7 in the retina, brain and somite boundaries. Loss of function of Chd7 resulted in several morphological defects similar to those observed in patients with CHARGE syndrome. We then performed a detailed analysis of uncharacterized defects of CHARGE syndrome and show that the presence of Chd7 is crucial for proper neural, retinal and vertebral development in zebrafish. These data provide new insights on the role of Chd7 and the mechanistic link between defects in the Chd7 gene and the organs and systems dysfunction associated with CHARGE syndrome. Furthermore, on the basis of the overlap in clinical features between zebrafish chd7 morphants and CHARGE syndrome, we suggest zebrafish can be a valuable in vivo tool to further understand the pathophysiological mechanisms underlying the abnormalities associated with CHARGE syndrome.

Materials and Methods

Animals

Wild-type AB, Isl1-GFP WIK transgenic and Fli1-GFP transgenic zebrafish (Danio rerio) embryos were raised at 28.5°C, and collected and staged using standard methods [64]. Wild-type AB stains fish and Fli1-GFP transgenic zebrafish were purchased from the Zebrafish International Resource Center (ZIRC; University of Oregon, Eugene, OR). Isl1-GFP WIK transgenic line was kindly provided by Dr. Hitoshi Okamoto (RIKEN Brain Science Institute, Wako, Japan) [65]. Embryos and larvae were anaesthetized in 0.02% tricaine (MS-222; Sigma Chemical, St. Louis, MO) in phosphate-buffered saline (PBS) prior to all procedures.

Ethics Statement

All protocols were carried out in compliance with the guidelines stipulated by the Canadian Council for Animal Care (CCAC), the CHU Sainte-Justine Research Center, as well as the Institutional Animal Care and Use Committee (IACUC) at Syracuse University and at the University of Montreal. This study was approved by the CHU Sainte-Justine Research Center ,University of Montreal (ZF-09-60/Category B) and the Syracuse University (IACUC # 07-010) ethics committees.

Design and synthesis of DIG-labeled RNA probes for chd7, efnb2a and ttna

Total RNA was isolated from blastula stage zebrafish embryos using the RNeasy Mini Kit (Quiagen, Valencia, CA). Reverse transcription was performed to make cDNA using the QuantiTect Reverse Transcription Kit (Quiagen, Valencia, CA). Chd7 fragments were amplified by PCR using 5′-GCTATTGACCGCTTCTCTCG-3′ and 5′-TGCTCCTTTACGCAGGAGAT-3′ primers (resulting in a 362 bp amplicon from about halfway through the cDNA sequence) and cloned into the pGEM-T Easy vector (Promega, Madison, WI) following the manufacturer's protocol. Zebrafish efnb2a [66] and ttna [67] plasmids were kind gifts from Dr. Steven Wilson and Dr. Andrew Oates respectively. To synthesize the antisense probe, the previously constructed plasmid was linearized using the restriction enzyme SpeI (New England BioLabs, Inc., Ipswich, MA) for 2 hours at 37°C. Linearization was confirmed by gel electrophoresis using a 1% agarose gel. Linearized plasmid was incubated with T7 RNA Polymerase (Roche Diagnostics, Indianapolis, IN) in the presence of digoxigenin (DIG) label (Roche Diagnostics, Indianapolis, IN) and RNase Inhibitor (Roche Diagnostics, Indianapolis, IN) for 2 hours at 37°C. To synthesize the sense strand, the restriction enzyme ApaI (New England Biolabs, Inc., Ipswich, MA) and SP6 RNA Polymerase (Roche Diagnostics, Indianapolis, IN) were used. Plasmid DNA was eliminated with DNase (Roche Diagnostics, Indianapolis, IN) by incubation at 37°C for 20 minutes. DIG-labeled RNA was precipitated in 0.2 M EDTA, 4 M LiCl, and 100% ethanol overnight at −20°C, and then resuspended in DEPC water. Agarose gel electrophoresis was used to confirm the presence of a purified probe and any unused probe was stored at −20°C.

Whole mount in situ hybridization analysis

Whole mount in situ hybridization was performed on staged zebrafish embryos using both sense and antisense chd7, efnb2a and ttna riboprobes. Methods for in situ hydridization analysis followed [68], [69]. Briefly, staged embryos were fixed overnight in 4% paraformaldehyde, and then dehydrated in methanol. When ready to use, embryos were rehydrated in phosphate buffered saline with 0.1% Tween-20 (PBSt). Embryos were permeabilized by proteinase K digestion and then hybridized with the riboprobes overnight at 70°C. The next day, embryos were put through graded solutions of 75%, 50%, and 25% prehybridized solution in 2× saline-sodium citrate (SSC) followed by a wash in 0.2× SSC for 30 minutes at 68°C. They were then placed in blocking solution for several hours, and incubated in α-DIG antibody overnight. Finally, embryos were washed again and incubated in staining solution in the dark until sufficient staining appeared on the embryos. Embryos were dehydrated in methanol to facilitate clearing of background staining and then rehydrated in PBSt. Embryos were stored in glycerol.

Embryos in glycerol were visualized using a Zeiss M2 Bio Stereomicroscope, with motorized focus drive and X-Cite UV light source with GFP filter. Images were captured using a Zeiss Axiocam digital camera connected to an Antec PC and processed with Adobe Photoshop 7.0.

Generation of zebrafish chd7 morpholino and mRNA rescue experiments

To eliminate Chd7 function, two types of antisense morpholino oligonucleotide were used to disrupt the translation of chd7 transcripts. The translation-blocking morpholino, 5′-TGCAGCCAAGCTTAGAAGCAGGAC-3′ and the splice blocking, 5′-TTATTTTCTGGCACTAACCATGTCC-3′ were synthesized by Gene Tools (Philomath, OR). The morpholino was injected into single-cell stage zebrafish embryos at doses of 2 ng/embryo, 4 ng/embryo and 6 ng/embryo. To validate the function of the chd7 splice blocking morpholino, RT-PCR was used to check for improper splicing using the following primer set: forward- 5′-AGGTGGACTCCGAAGGAAAC-3′, reverse- 5′-CCGTCATCACCACATTTGAG-3′. Amplified cDNA was visualized using gel electrophoresis.

To confirm the specificity of the chd7 morpholinos, a mismatch morpholino (lower case) was used: TGgAcCCAAcCTTAcAAcCAGGAC (Gene Tools. OR). Furthermore, rescue experiments were performed. Full-length wild type zebrafish chd7 gene was subcloned into pcs2+ vector and capped mRNA was synthesized using the Sp6 promoter and the mMessage mMachine Kit (Ambion). Three hundred picograms of the synthetic mRNA was then injected into embryos at the one-cell stage. Rescued embryos were visualized and counted using the Olympus SZX12 stereoscope.

Injected and uninjected embryos were then incubated in embryo media at 28.5°C for 24 h, after which they were assessed for viability. Wild type, Isl1-GFP WIK and Fli1-GFP transgenic zebrafish embryos injected with chd7 morpholino were assessed for morphological differences from control morpholino injected or uninjected embryos under an Olympus SZX12 stereoscope at 24, 48 and 72 hpf. Isl1-GFP transgenic zebrafish were stage matched to 48 hpf and fixed in 4% paraformaldehyde for 4 h and then analyzed for differences in branchiomotor neuron development and migration from uninjected controls using a Zeiss AxioImager Z1 compound microscope. Images were photographed using a Zeiss LSM 510 confocal microscope at 488 nm under a 20× objective, and were compiled using Zeiss LSM Image Browser software. Fli1-GPF zebrafish were staged to 34–36 hpf and screened for CNC segmentation defects using an X-Cite UV light source with GFP filter mounted to a Zeiss M2 Bio Stereomicroscope.

Immunohistological procedures

Zebrafish specimens were fixed in 4% paraformaldehyde and were either used for whole-mount immunostaining or embedded in paraffin. Transverse sections (1.5–3 µm) of paraffin-embedded specimens were deparaffinized in xylene and were rehydrated in a graded series of ethanol. Serial sections were collected from the central retina.

For immunofluorescence microscopy, whole embryos and eye sections were washed several times in PBS and permeabilized for 30 min in 4% Triton-X 100 containing 2% bovine serum albumin (BSA) and 10% goat serum. Following permeabilization, tissues and retinal sections were incubated in the primary antibody 3A10 (Developmental Studies Hybridoma Bank; 1∶500) for 48 h at 4°C on a shaker. Tissues and eye sections were washed several times in PBS over a 24 h period, and then incubated in the secondary antibody conjugated with Alexa Fluor 488 (Molecular Probes, Carlsbad, CA, 1∶2000) for 4–6 h at room temperature. Animals were washed in PBS several times, de-yolked, cleared in 70% glycerol and mounted. Z-stack images were photographed using a Zeiss LSM 510 confocal microscope under a 20× objective, and were compiled using Zeiss LSM Image Browser software.

For immunoperoxidase methods, slides were incubated with a blocking serum (Vectastain; Vector Laboratories, Burlingame, CA) for 45 minutes, after which they were blotted and then overlaid with the primary antibodies Chd7 (Santa Cruz Biotechnology, Santa Cruz, CA ; 1/600) and zn-8 (Developmental Studies Hybridoma Bank; 1/200) for 18 hours at 4°C. The slides were washed 3 times in PBS, pH 7.4, and incubated with secondary antibodies (anti- mouse (1∶1000) or anti- rabbit (1∶1000), Vectastain) for 1 hour at room temperature, followed by staining according to the avidin-biotin-peroxidase complex method (Vectastain ABC assay). Color was developed with 3,3-diaminobenzidine (Dako Diagnostics Inc., Mississauga, ON, Canada) containing hydrogen peroxide. Slides were counterstained with Harris modified hematoxylin (Fisher Scientific, Ottawa, ON, Canada). Eye sections were also stained with 1% Toluidine Blue. Images were captured using a Leica DMR microscope mounted with a Qimaging Retiga 1300 camera.

Skeletal staining

Juvenile zebrafish were fixed in 4% paraformaldehyde in PBS for 48 hr at 4°C and dehydrated to 100% ethanol over 2 days. Fish were then cleared in a 1% KOH solution containing approximately 4.5 ml of 3% H₂O₂ per 100 ml 1% KOH for 2 h, followed by 2 h in half this concentration (2.25 ml H₂O₂ per 100 ml 1% KOH). This latter step ensures a stepwise progression from H₂O₂ to water, which helps reduce tissue expansion. Zebrafish were then rinsed twice in distilled water, and transferred to a 30% saturated sodium tetraborate solution (Borax; Acros Organics) in water, overnight. Any remaining soft tissues were digested in a solution containing 1.0% trypsin and 2.0% Borax in water for 2–4 h. Digestion was deemed complete when the primary axis of the axial skeleton became visible. The fish were subsequently rinsed three times in distilled water to remove all residual trypsin, prior to staining for 12 h or more in a solution of 75% ethanol and Alizarin red S. Vertebral mineralization was assessed after Alizarin red S staining. Fish were examined using a Leica M205FA stereomicroscope. Digital images were collected with Leica DFC490 camera.

Calcein staining was performed as described [70]. Briefly, fish were anesthetized in 0.6 mM MS-222 buffered to pH 7.0 and immersed in 0.2% calcein in 10% Hank's solution buffered to pH 7.2 for 10 min followed by three 10 min washes in Hank's solution. Vertebral mineralization was assessed after calcein staining using a Leica DMR microscope. Digital images were collected with a Qimaging Retiga 1300 camera.

Sample number and statistical analysis

The sample number (n) refers to the batch number and each batch number consisted of at least 40 fish. For each data set, we provided the batch number followed by the total fish number used. The sample number for wild-type fish consisted of 12 batches with a total of 510 fish used (n = 12,510 fish); 12 batches with a total of 436 fish for chd7 morphants (n = 12,436 fish); 12 batches with a total of 498 fish for control-MO injected embryos (n = 12, 498 fish) and 6 batches with a total of 249 mRNA rescue experiments (n = 6, 249 fish). Immunohistochemistry were performed on at least 6 batches for both control-MO and chd7-MO fish. Statistical analyses were performed and data were plotted in SigmaPlot 11.0 (Systat Software Inc., San Jose, CA, USA). Significance was determined using paired Student's t-tests, one-way anovas and Fisher's least significant difference tests for normally distributed, equal variance data. Kruskal–Wallis anova and Dunn's method of comparison were used for non-normal distributions.

Supporting Information

[Figure omitted. See PDF.]

Figure S1.

Chd7- MO injections are dosage dependent. As the concentration of the chd7-MO injection increased, the phenotypic defects became more severe. (A) Control-MO injected zebrafish showed no phenotypic defects 48 hpf and were comparable to wild type zebrafish at the same age. Embryos injected with 2 ng/nl (B), 4 ng/nl (C), or 6 ng/nl (D) chd7-MO showed increasing severity of developmental defects with increasing MO concentration.

https://doi.org/10.1371/journal.pone.0031650.s001

(TIF)

Figure S2.

Chd7 plays an essential role in photoreceptor development. The photoreceptor layer of control-MO (A) and chd7-MO-injected (B) embryos were stained with Zpr-1. Chd7 morphants lacked the photoreceptor layer.

https://doi.org/10.1371/journal.pone.0031650.s002

(TIF)

Table S1.

Measurement of zebrafish size at different ages.

https://doi.org/10.1371/journal.pone.0031650.s003

(TIF)

Acknowledgments

We are grateful to G. Laliberté and R. Milette for animal care. We would also like to thank Jeff Amack for the use of microinjection equipment and for Fli1-GFP transgenic fish. Thanks to Fiona Foley for help with the microinjections. The monoclonal antibodies 3A10 and zn-8 developed by Thomas M. Jessell and Jane Dodd were obtained from the Developmental Studies Hybridoma Bank, developed under the auspices of the National Institute of Child Health and Human Development, and maintained by the University of Iowa, Department of Biological Sciences (Iowa City, IA).

Author Contributions

Conceived and designed the experiments: SP NJM RCA FM. Performed the experiments: SP NJM CZ. Analyzed the data: SP NJM. Contributed reagents/materials/analysis tools: PD FM RCA. Wrote the paper: SP NJM. Read and edited the manuscript: SP NJM FM PD RCA.

Citation: Patten SA, Jacobs-McDaniels NL, Zaouter C, Drapeau P, Albertson RC, Moldovan F (2012) Role of Chd7 in Zebrafish: A Model for CHARGE Syndrome. PLoS ONE 7(2): e31650. https://doi.org/10.1371/journal.pone.0031650

References

1. Vissers LE, van Ravenswaaij CM, Admiraal R, Hurst JA, de Vries BB, et al. (2004) Mutations in a new member of the chromodomain gene family cause CHARGE syndrome. Nat Genet 36: 955–957.LE VissersCM van RavenswaaijR. AdmiraalJA HurstBB de Vries2004Mutations in a new member of the chromodomain gene family cause CHARGE syndrome.Nat Genet36955957

2. Aramaki M, Udaka T, Kosaki R, Makita Y, Okamoto N, et al. (2006) Phenotypic spectrum of CHARGE syndrome with CHD7 mutations. J Pediatr 148: 410–414.M. AramakiT. UdakaR. KosakiY. MakitaN. Okamoto2006Phenotypic spectrum of CHARGE syndrome with CHD7 mutations.J Pediatr148410414

3. Lalani SR, Safiullah AM, Fernbach SD, Harutyunyan KG, Thaller C, et al. (2006) Spectrum of CHD7 mutations in 110 individuals with CHARGE syndrome and genotype-phenotype correlation. Am J Hum Genet 78: 303–314.SR LalaniAM SafiullahSD FernbachKG HarutyunyanC. Thaller2006Spectrum of CHD7 mutations in 110 individuals with CHARGE syndrome and genotype-phenotype correlation.Am J Hum Genet78303314

4. Sanlaville D, Etchevers HC, Gonzales M, Martinovic J, Clement-Ziza M, et al. (2006) Phenotypic spectrum of CHARGE syndrome in fetuses with CHD7 truncating mutations correlates with expression during human development. J Med Genet 43: 211–217.D. SanlavilleHC EtcheversM. GonzalesJ. MartinovicM. Clement-Ziza2006Phenotypic spectrum of CHARGE syndrome in fetuses with CHD7 truncating mutations correlates with expression during human development.J Med Genet43211217

5. Jongmans MC, Admiraal RJ, van der Donk KP, Vissers LE, Baas AF, et al. (2006) CHARGE syndrome: the phenotypic spectrum of mutations in the CHD7 gene. J Med Genet 43: 306–314.MC JongmansRJ AdmiraalKP van der DonkLE VissersAF Baas2006CHARGE syndrome: the phenotypic spectrum of mutations in the CHD7 gene.J Med Genet43306314

6. Lalani SR, Safiullah AM, Fernbach SD, Phillips M, Bacino CA, et al. (2005) SNP genotyping to screen for a common deletion in CHARGE syndrome. BMC Med Genet 6: 8.SR LalaniAM SafiullahSD FernbachM. PhillipsCA Bacino2005SNP genotyping to screen for a common deletion in CHARGE syndrome.BMC Med Genet68

7. Wincent J, Schulze A, Schoumans J (2009) Detection of CHD7 deletions by MLPA in CHARGE syndrome patients with a less typical phenotype. Eur J Med Genet 52: 271–272.J. WincentA. SchulzeJ. Schoumans2009Detection of CHD7 deletions by MLPA in CHARGE syndrome patients with a less typical phenotype.Eur J Med Genet52271272

8. Vuorela P, Ala-Mello S, Saloranta C, Penttinen M, Poyhonen M, et al. (2007) Molecular analysis of the CHD7 gene in CHARGE syndrome: identification of 22 novel mutations and evidence for a low contribution of large CHD7 deletions. Genet Med 9: 690–694.P. VuorelaS. Ala-MelloC. SalorantaM. PenttinenM. Poyhonen2007Molecular analysis of the CHD7 gene in CHARGE syndrome: identification of 22 novel mutations and evidence for a low contribution of large CHD7 deletions.Genet Med9690694

9. Wessels K, Bohnhorst B, Luhmer I, Morlot S, Bohring A, et al. (2010) Novel CHD7 mutations contributing to the mutation spectrum in patients with CHARGE syndrome. Eur J Med Genet 53: 280–285.K. WesselsB. BohnhorstI. LuhmerS. MorlotA. Bohring2010Novel CHD7 mutations contributing to the mutation spectrum in patients with CHARGE syndrome.Eur J Med Genet53280285

10. Carey JC (2005) CHARGE syndrome: 2005. Am J Med Genet A 133A: 227.JC Carey2005CHARGE syndrome: 2005.Am J Med Genet A133A227

11. Blake KD, Prasad C (2006) CHARGE syndrome. Orphanet J Rare Dis 1: 34.KD BlakeC. Prasad2006CHARGE syndrome.Orphanet J Rare Dis134

12. Marfella CG, Imbalzano AN (2007) The Chd family of chromatin remodelers. Mutat Res 618: 30–40.CG MarfellaAN Imbalzano2007The Chd family of chromatin remodelers.Mutat Res6183040

13. Allen MD, Religa TL, Freund SM, Bycroft M (2007) Solution structure of the BRK domains from CHD7. J Mol Biol 371: 1135–1140.MD AllenTL ReligaSM FreundM. Bycroft2007Solution structure of the BRK domains from CHD7.J Mol Biol37111351140

14. Schnetz MP, Bartels CF, Shastri K, Balasubramanian D, Zentner GE, et al. (2009) Genomic distribution of CHD7 on chromatin tracks H3K4 methylation patterns. Genome Res 19: 590–601.MP SchnetzCF BartelsK. ShastriD. BalasubramanianGE Zentner2009Genomic distribution of CHD7 on chromatin tracks H3K4 methylation patterns.Genome Res19590601

15. Srinivasan S, Dorighi KM, Tamkun JW (2008) Drosophila Kismet regulates histone H3 lysine 27 methylation and early elongation by RNA polymerase II. PLoS Genet 4: e1000217.S. SrinivasanKM DorighiJW Tamkun2008Drosophila Kismet regulates histone H3 lysine 27 methylation and early elongation by RNA polymerase II.PLoS Genet4e1000217

16. Takada I, Mihara M, Suzawa M, Ohtake F, Kobayashi S, et al. (2007) A histone lysine methyltransferase activated by non-canonical Wnt signalling suppresses PPAR-gamma transactivation. Nat Cell Biol 9: 1273–1285.I. TakadaM. MiharaM. SuzawaF. OhtakeS. Kobayashi2007A histone lysine methyltransferase activated by non-canonical Wnt signalling suppresses PPAR-gamma transactivation.Nat Cell Biol912731285

17. Bajpai R, Chen DA, Rada-Iglesias A, Zhang J, Xiong Y, et al. (2010) CHD7 cooperates with PBAF to control multipotent neural crest formation. Nature 463: 958–962.R. BajpaiDA ChenA. Rada-IglesiasJ. ZhangY. Xiong2010CHD7 cooperates with PBAF to control multipotent neural crest formation.Nature463958962

18. Bosman EA, Penn AC, Ambrose JC, Kettleborough R, Stemple DL, et al. (2005) Multiple mutations in mouse Chd7 provide models for CHARGE syndrome. Hum Mol Genet 14: 3463–3476.EA BosmanAC PennJC AmbroseR. KettleboroughDL Stemple2005Multiple mutations in mouse Chd7 provide models for CHARGE syndrome.Hum Mol Genet1434633476

19. Jacobs-McDaniels NL, Albertson RC (2011) Chd7 plays a critical role in controlling left-right symmetry during zebrafish somitogenesis. Dev Dyn 240: 2272–2280.NL Jacobs-McDanielsRC Albertson2011Chd7 plays a critical role in controlling left-right symmetry during zebrafish somitogenesis.Dev Dyn24022722280

20. Gao X, Gordon D, Zhang D, Browne R, Helms C, et al. (2007) CHD7 gene polymorphisms are associated with susceptibility to idiopathic scoliosis. Am J Hum Genet 80: 957–965.X. GaoD. GordonD. ZhangR. BrowneC. Helms2007CHD7 gene polymorphisms are associated with susceptibility to idiopathic scoliosis.Am J Hum Genet80957965

21. Gregory CA, Gunn WG, Peister A, Prockop DJ (2004) An Alizarin red-based assay of mineralization by adherent cells in culture: comparison with cetylpyridinium chloride extraction. Anal Biochem 329: 77–84.CA GregoryWG GunnA. PeisterDJ Prockop2004An Alizarin red-based assay of mineralization by adherent cells in culture: comparison with cetylpyridinium chloride extraction.Anal Biochem3297784

22. Nasevicius A, Ekker SC (2000) Effective targeted gene ‘knockdown’ in zebrafish. Nat Genet 26: 216–220.A. NaseviciusSC Ekker2000Effective targeted gene ‘knockdown’ in zebrafish.Nat Genet26216220

23. Takahashi Y, Koizumi K, Takagi A, Kitajima S, Inoue T, et al. (2000) Mesp2 initiates somite segmentation through the Notch signalling pathway. Nat Genet 25: 390–396.Y. TakahashiK. KoizumiA. TakagiS. KitajimaT. Inoue2000Mesp2 initiates somite segmentation through the Notch signalling pathway.Nat Genet25390396

24. Koizumi K, Nakajima M, Yuasa S, Saga Y, Sakai T, et al. (2001) The role of presenilin 1 during somite segmentation. Development 128: 1391–1402.K. KoizumiM. NakajimaS. YuasaY. SagaT. Sakai2001The role of presenilin 1 during somite segmentation.Development12813911402

25. Shaw KM, Castranova DA, Pham VN, Kamei M, Kidd KR, et al. (2006) fused-somites-like mutants exhibit defects in trunk vessel patterning. Dev Dyn 235: 1753–1760.KM ShawDA CastranovaVN PhamM. KameiKR Kidd2006fused-somites-like mutants exhibit defects in trunk vessel patterning.Dev Dyn23517531760

26. Childs S, Chen JN, Garrity DM, Fishman MC (2002) Patterning of angiogenesis in the zebrafish embryo. Development 129: 973–982.S. ChildsJN ChenDM GarrityMC Fishman2002Patterning of angiogenesis in the zebrafish embryo.Development129973982

27. Hurd EA, Poucher HK, Cheng K, Raphael Y, Martin DM (2010) The ATP-dependent chromatin remodeling enzyme CHD7 regulates pro-neural gene expression and neurogenesis in the inner ear. Development 137: 3139–3150.EA HurdHK PoucherK. ChengY. RaphaelDM Martin2010The ATP-dependent chromatin remodeling enzyme CHD7 regulates pro-neural gene expression and neurogenesis in the inner ear.Development13731393150

28. Aramaki M, Kimura T, Udaka T, Kosaki R, Mitsuhashi T, et al. (2007) Embryonic expression profile of chicken CHD7, the ortholog of the causative gene for CHARGE syndrome. Birth Defects Res A Clin Mol Teratol 79: 50–57.M. AramakiT. KimuraT. UdakaR. KosakiT. Mitsuhashi2007Embryonic expression profile of chicken CHD7, the ortholog of the causative gene for CHARGE syndrome.Birth Defects Res A Clin Mol Teratol795057

29. Melicharek DJ, Ramirez LC, Singh S, Thompson R, Marenda DR (2010) Kismet/CHD7 regulates axon morphology, memory and locomotion in a Drosophila model of CHARGE syndrome. Hum Mol Genet 19: 4253–4264.DJ MelicharekLC RamirezS. SinghR. ThompsonDR Marenda2010Kismet/CHD7 regulates axon morphology, memory and locomotion in a Drosophila model of CHARGE syndrome.Hum Mol Genet1942534264

30. Christiansen HE, Lang MR, Pace JM, Parichy DM (2009) Critical early roles for col27a1a and col27a1b in zebrafish notochord morphogenesis, vertebral mineralization and post-embryonic axial growth. PLoS One 4: e8481.HE ChristiansenMR LangJM PaceDM Parichy2009Critical early roles for col27a1a and col27a1b in zebrafish notochord morphogenesis, vertebral mineralization and post-embryonic axial growth.PLoS One4e8481

31. Smith A, Avaron F, Guay D, Padhi BK, Akimenko MA (2006) Inhibition of BMP signaling during zebrafish fin regeneration disrupts fin growth and scleroblasts differentiation and function. Dev Biol 299: 438–454.A. SmithF. AvaronD. GuayBK PadhiMA Akimenko2006Inhibition of BMP signaling during zebrafish fin regeneration disrupts fin growth and scleroblasts differentiation and function.Dev Biol299438454

32. Layman WS, McEwen DP, Beyer LA, Lalani SR, Fernbach SD, et al. (2009) Defects in neural stem cell proliferation and olfaction in Chd7 deficient mice indicate a mechanism for hyposmia in human CHARGE syndrome. Hum Mol Genet 18: 1909–1923.WS LaymanDP McEwenLA BeyerSR LalaniSD Fernbach2009Defects in neural stem cell proliferation and olfaction in Chd7 deficient mice indicate a mechanism for hyposmia in human CHARGE syndrome.Hum Mol Genet1819091923

33. Akisu M, Ozkinay F, Ozyurek R, Kucuktas A, Kultursay N (1998) The CHARGE association in a newborn infant. Turk J Pediatr 40: 283–287.M. AkisuF. OzkinayR. OzyurekA. KucuktasN. Kultursay1998The CHARGE association in a newborn infant.Turk J Pediatr40283287

34. Ben Becher S, Ganouni S, Cheour M, Bouaziz A, Boudhina T (1994) [CHARGE association]. Arch Pediatr 1: 1115–1117.S. Ben BecherS. GanouniM. CheourA. BouazizT. Boudhina1994[CHARGE association].Arch Pediatr111151117

35. Van Meter TD, Weaver DD (1996) Oculo-auriculo-vertebral spectrum and the CHARGE association: clinical evidence for a common pathogenetic mechanism. Clin Dysmorphol 5: 187–196.TD Van MeterDD Weaver1996Oculo-auriculo-vertebral spectrum and the CHARGE association: clinical evidence for a common pathogenetic mechanism.Clin Dysmorphol5187196

36. Larison KD, Bremiller R (1990) Early onset of phenotype and cell patterning in the embryonic zebrafish retina. Development 109: 567–576.KD LarisonR. Bremiller1990Early onset of phenotype and cell patterning in the embryonic zebrafish retina.Development109567576

37. Link BA, Fadool JM, Malicki J, Dowling JE (2000) The zebrafish young mutation acts non-cell-autonomously to uncouple differentiation from specification for all retinal cells. Development 127: 2177–2188.BA LinkJM FadoolJ. MalickiJE Dowling2000The zebrafish young mutation acts non-cell-autonomously to uncouple differentiation from specification for all retinal cells.Development12721772188

38. Masai I, Lele Z, Yamaguchi M, Komori A, Nakata A, et al. (2003) N-cadherin mediates retinal lamination, maintenance of forebrain compartments and patterning of retinal neurites. Development 130: 2479–2494.I. MasaiZ. LeleM. YamaguchiA. KomoriA. Nakata2003N-cadherin mediates retinal lamination, maintenance of forebrain compartments and patterning of retinal neurites.Development13024792494

39. Erdmann B, Kirsch FP, Rathjen FG, More MI (2003) N-cadherin is essential for retinal lamination in the zebrafish. Dev Dyn 226: 570–577.B. ErdmannFP KirschFG RathjenMI More2003N-cadherin is essential for retinal lamination in the zebrafish.Dev Dyn226570577

40. Schmitt EA, Dowling JE (1999) Early retinal development in the zebrafish, Danio rerio: light and electron microscopic analyses. J Comp Neurol 404: 515–536.EA SchmittJE Dowling1999Early retinal development in the zebrafish, Danio rerio: light and electron microscopic analyses.J Comp Neurol404515536

41. Melicharek D, Shah A, DiStefano G, Gangemi AJ, Orapallo A, et al. (2008) Identification of novel regulators of atonal expression in the developing Drosophila retina. Genetics 180: 2095–2110.D. MelicharekA. ShahG. DiStefanoAJ GangemiA. Orapallo2008Identification of novel regulators of atonal expression in the developing Drosophila retina.Genetics18020952110

42. Jarman AP, Sun Y, Jan LY, Jan YN (1995) Role of the proneural gene, atonal, in formation of Drosophila chordotonal organs and photoreceptors. Development 121: 2019–2030.AP JarmanY. SunLY JanYN Jan1995Role of the proneural gene, atonal, in formation of Drosophila chordotonal organs and photoreceptors.Development12120192030

43. Yokoi H, Yan YL, Miller MR, BreMiller RA, Catchen JM, et al. (2009) Expression profiling of zebrafish sox9 mutants reveals that Sox9 is required for retinal differentiation. Dev Biol 329: 1–15.H. YokoiYL YanMR MillerRA BreMillerJM Catchen2009Expression profiling of zebrafish sox9 mutants reveals that Sox9 is required for retinal differentiation.Dev Biol329115

44. Lacombe D (1994) Facial palsy and cranial nerve abnormalities in CHARGE association. Am J Med Genet 49: 351–353.D. Lacombe1994Facial palsy and cranial nerve abnormalities in CHARGE association.Am J Med Genet49351353

45. Byerly KA, Pauli RM (1993) Cranial nerve abnormalities in CHARGE association. Am J Med Genet 45: 751–757.KA ByerlyRM Pauli1993Cranial nerve abnormalities in CHARGE association.Am J Med Genet45751757

46. Huang X, Saint-Jeannet JP (2004) Induction of the neural crest and the opportunities of life on the edge. Dev Biol 275: 1–11.X. HuangJP Saint-Jeannet2004Induction of the neural crest and the opportunities of life on the edge.Dev Biol275111

47. Grenier J, Teillet MA, Grifone R, Kelly RG, Duprez D (2009) Relationship between neural crest cells and cranial mesoderm during head muscle development. PLoS One 4: e4381.J. GrenierMA TeilletR. GrifoneRG KellyD. Duprez2009Relationship between neural crest cells and cranial mesoderm during head muscle development.PLoS One4e4381

48. Lwigale PY, Bronner-Fraser M (2009) Semaphorin3A/neuropilin-1 signaling acts as a molecular switch regulating neural crest migration during cornea development. Dev Biol 336: 257–265.PY LwigaleM. Bronner-Fraser2009Semaphorin3A/neuropilin-1 signaling acts as a molecular switch regulating neural crest migration during cornea development.Dev Biol336257265

49. Knight RD, Schilling TF (2006) Cranial neural crest and development of the head skeleton. Adv Exp Med Biol 589: 120–133.RD KnightTF Schilling2006Cranial neural crest and development of the head skeleton.Adv Exp Med Biol589120133

50. Amiel J, Attiee-Bitach T, Marianowski R, Cormier-Daire V, Abadie V, et al. (2001) Temporal bone anomaly proposed as a major criteria for diagnosis of CHARGE syndrome. Am J Med Genet 99: 124–127.J. AmielT. Attiee-BitachR. MarianowskiV. Cormier-DaireV. Abadie2001Temporal bone anomaly proposed as a major criteria for diagnosis of CHARGE syndrome.Am J Med Genet99124127

51. Blake KD, Davenport SL, Hall BD, Hefner MA, Pagon RA, et al. (1998) CHARGE association: an update and review for the primary pediatrician. Clin Pediatr (Phila) 37: 159–173.KD BlakeSL DavenportBD HallMA HefnerRA Pagon1998CHARGE association: an update and review for the primary pediatrician.Clin Pediatr (Phila)37159173

52. Morgan D, Bailey M, Phelps P, Bellman S, Grace A, et al. (1993) Ear-nose-throat abnormalities in the CHARGE association. Arch Otolaryngol Head Neck Surg 119: 49–54.D. MorganM. BaileyP. PhelpsS. BellmanA. Grace1993Ear-nose-throat abnormalities in the CHARGE association.Arch Otolaryngol Head Neck Surg1194954

53. Abadie V, Wiener-Vacher S, Morisseau-Durand MP, Poree C, Amiel J, et al. (2000) Vestibular anomalies in CHARGE syndrome: investigations on and consequences for postural development. Eur J Pediatr 159: 569–574.V. AbadieS. Wiener-VacherMP Morisseau-DurandC. PoreeJ. Amiel2000Vestibular anomalies in CHARGE syndrome: investigations on and consequences for postural development.Eur J Pediatr159569574

54. Satar B, Mukherji SK, Telian SA (2003) Congenital aplasia of the semicircular canals. Otol Neurotol 24: 437–446.B. SatarSK MukherjiSA Telian2003Congenital aplasia of the semicircular canals.Otol Neurotol24437446

55. Tellier AL, Cormier-Daire V, Abadie V, Amiel J, Sigaudy S, et al. (1998) CHARGE syndrome: report of 47 cases and review. Am J Med Genet 76: 402–409.AL TellierV. Cormier-DaireV. AbadieJ. AmielS. Sigaudy1998CHARGE syndrome: report of 47 cases and review.Am J Med Genet76402409

56. Souriau J, Gimenes M, Blouin C, Benbrik I, Benbrik E, et al. (2005) CHARGE syndrome: developmental and behavioral data. Am J Med Genet A 133A: 278–281.J. SouriauM. GimenesC. BlouinI. BenbrikE. Benbrik2005CHARGE syndrome: developmental and behavioral data.Am J Med Genet A133A278281

57. Schimmang T (2007) Expression and functions of FGF ligands during early otic development. Int J Dev Biol 51: 473–481.T. Schimmang2007Expression and functions of FGF ligands during early otic development.Int J Dev Biol51473481

58. Chang W, Brigande JV, Fekete DM, Wu DK (2004) The development of semicircular canals in the inner ear: role of FGFs in sensory cristae. Development 131: 4201–4211.W. ChangJV BrigandeDM FeketeDK Wu2004The development of semicircular canals in the inner ear: role of FGFs in sensory cristae.Development13142014211

59. Alvarez Y, Alonso MT, Vendrell V, Zelarayan LC, Chamero P, et al. (2003) Requirements for FGF3 and FGF10 during inner ear formation. Development 130: 6329–6338.Y. AlvarezMT AlonsoV. VendrellLC ZelarayanP. Chamero2003Requirements for FGF3 and FGF10 during inner ear formation.Development13063296338

60. Hammond KL, Whitfield TT (2006) The developing lamprey ear closely resembles the zebrafish otic vesicle: otx1 expression can account for all major patterning differences. Development 133: 1347–1357.KL HammondTT Whitfield2006The developing lamprey ear closely resembles the zebrafish otic vesicle: otx1 expression can account for all major patterning differences.Development13313471357

61. Thomas IT, Frias JL (1987) The heart in selected congenital malformations. A lesson in pathogenetic relationships. Ann Clin Lab Sci 17: 207–210.IT ThomasJL Frias1987The heart in selected congenital malformations. A lesson in pathogenetic relationships.Ann Clin Lab Sci17207210

62. Jyonouchi S, McDonald-McGinn DM, Bale S, Zackai EH, Sullivan KE (2009) CHARGE (coloboma, heart defect, atresia choanae, retarded growth and development, genital hypoplasia, ear anomalies/deafness) syndrome and chromosome 22q11.2 deletion syndrome: a comparison of immunologic and nonimmunologic phenotypic features. Pediatrics 123: e871–877.S. JyonouchiDM McDonald-McGinnS. BaleEH ZackaiKE Sullivan2009CHARGE (coloboma, heart defect, atresia choanae, retarded growth and development, genital hypoplasia, ear anomalies/deafness) syndrome and chromosome 22q11.2 deletion syndrome: a comparison of immunologic and nonimmunologic phenotypic features.Pediatrics123e871877

63. Devriendt K, Swillen A, Fryns JP (1998) Deletion in chromosome region 22q11 in a child with CHARGE association. Clin Genet 53: 408–410.K. DevriendtA. SwillenJP Fryns1998Deletion in chromosome region 22q11 in a child with CHARGE association.Clin Genet53408410

64. Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF (1995) Stages of embryonic development of the zebrafish. Dev Dyn 203: 253–310.CB KimmelWW BallardSR KimmelB. UllmannTF Schilling1995Stages of embryonic development of the zebrafish.Dev Dyn203253310

65. Uemura O, Okada Y, Ando H, Guedj M, Higashijima S, et al. (2005) Comparative functional genomics revealed conservation and diversification of three enhancers of the isl1 gene for motor and sensory neuron-specific expression. Dev Biol 278: 587–606.O. UemuraY. OkadaH. AndoM. GuedjS. Higashijima2005Comparative functional genomics revealed conservation and diversification of three enhancers of the isl1 gene for motor and sensory neuron-specific expression.Dev Biol278587606

66. Barrios A, Poole RJ, Durbin L, Brennan C, Holder N, et al. (2003) Eph/Ephrin signaling regulates the mesenchymal-to-epithelial transition of the paraxial mesoderm during somite morphogenesis. Curr Biol 13: 1571–1582.A. BarriosRJ PooleL. DurbinC. BrennanN. Holder2003Eph/Ephrin signaling regulates the mesenchymal-to-epithelial transition of the paraxial mesoderm during somite morphogenesis.Curr Biol1315711582

67. Oates AC, Mueller C, Ho RK (2005) Cooperative function of deltaC and her7 in anterior segment formation. Dev Biol 280: 133–149.AC OatesC. MuellerRK Ho2005Cooperative function of deltaC and her7 in anterior segment formation.Dev Biol280133149

68. Albertson RC, Yelick PC (2005) Roles for fgf8 signaling in left-right patterning of the visceral organs and craniofacial skeleton. Dev Biol 283: 310–321.RC AlbertsonPC Yelick2005Roles for fgf8 signaling in left-right patterning of the visceral organs and craniofacial skeleton.Dev Biol283310321

69. Jacobs NL, Albertson RC, Wiles JR (2011) Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology. J Vis Exp. NL JacobsRC AlbertsonJR Wiles2011Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology.J Vis Exp

70. Du SJ, Frenkel V, Kindschi G, Zohar Y (2001) Visualizing normal and defective bone development in zebrafish embryos using the fluorescent chromophore calcein. Dev Biol 238: 239–246.SJ DuV. FrenkelG. KindschiY. Zohar2001Visualizing normal and defective bone development in zebrafish embryos using the fluorescent chromophore calcein.Dev Biol238239246