Rapgef1 paralog-mediated regulation of Wnt/β-catenin signaling orchestrates early embryo tissue patterning and morphogenesis

Abstract

RAPGEF1, a guanine nucleotide exchange factor, regulates signaling and cytoskeletal dynamics in mammalian cells, yet its role in development remains unclear as Rapgef1 null mouse embryos do not survive beyond implantation. We demonstrate that zebrafish rapgef1 is maternally expressed, and its paralogs, rapgef1a and rapgef1b, exhibit tissue and developmental stage-specific splicing. Disruption of rapgef1b caused brain and somite defects, impaired cranial neural crest specification, and microcephaly-like phenotypes, uncovering its previously uncharacterized functions in morphogenesis and tissue patterning. Transcriptomic analyses and differential gene expression provide fresh insights into the developmental functions of rapgef1b in presomitic mesoderm and somitogenesis by modulating the Wnt/β catenin signaling. Rapgef1b deficient embryos also showed spindle pole disorganization and chromosome mis-congression, linking Rapgef1 to centrosome-mediated mitotic fidelity. Together, our findings identify Rapgef1b as a key regulator of neural crest development, mesodermal morphogenesis, and early mitoses, highlighting its tissue-specific functions during vertebrate embryogenesis.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12964-026-02905-0.

Teaser

In this study, we show that rapgef1b is essential for shaping the embryonic brain, somites, and body axis by regulating gene expression, cell division, survival, and differentiation. Our findings reveal a new dimension of signaling-mediated cell fate specification during early vertebrate development.

Introduction

Guanine Nucleotide Exchange Factors (GEFs) serve as key signaling centers that activate small GTPases, enabling multiple effector functions to regulate adhesion, cytoskeletal dynamics, and signaling. One of the GEFs, RAPGEF1 (C3G), is predominantly expressed as a 140kD protein with a modular architecture consisting of a C-terminal catalytic domain, a central protein interaction region enriched in proline-rich Crk binding motifs (CBR), and an N-terminal E-cadherin binding domain [1]. RAPGEF1 localizes to the subcortical cytoskeleton, Golgi, nuclear speckles, centrosome, and exhibits nucleo-cytoplasmic exchange [2–5]. Its activity is tightly controlled through intramolecular interactions, post-translational modifications such as phosphorylation, and intermolecular protein interactions. RAPGEF1 functions as an enzyme and a scaffold by engaging with diverse signaling proteins [3, 6]. At the molecular level, it interacts with GRB2, CRK, CAS, Src family kinases, β-catenin, GSK3β, and cenexin in mammalian cells, regulating chromatin organization, splicing, cytoskeletal remodeling, and transcription [2, 5, 7, 8]. Deregulated RAPGEF1 levels and missense mutations are associated with multiple human pathologies, including cancers and neurological disorders [9–17]. Although tissue-specific, alternately spliced isoforms of Rapgef1 have been described in mice, their functions remain unknown [18, 19]. Notably, a novel brain-specific, long isoform of Rapgef1 has recently been identified in adult murine brains and human brain organoids [7, 18–21].

The loss-of-function studies underscore the developmental importance of Rapgef1. Mouse embryonic stem cells lacking Rapgef1 showed enhanced self-renewal but failed to undergo lineage differentiation [22, 23]. Further, Rapgef1 knockout mice embryos failed to survive beyond implantation, highlighting its essential role in early embryogenesis [24, 25]. Hypomorphic mice expressing a human Rapgef1 allele exhibited severe defects in cell adhesion, vasculogenesis, and differentiation [26–28]. However, the role of rapgef1 in early embryonic events such as cell fate specification, lineage determination, tissue patterning and morphogenesis remains unresolved.

To address this gap, we investigated rapgef1 function in zebrafish, which possess two paralogs, rapgef1a and rapgef1b. Knockdown of rapgef1a has been reported to cause pan developmental defects, including brain and blood vessel anomalies, motor neuron defects, and impaired locomotor activity [13]. In contrast, the function of the other homolog, rapgef1b, has not been explored. Here, we examined the spatiotemporal expression of transcripts generated from the two paralogs and demonstrate that rapgef1a and rapgef1b are both expressed in early embryogenesis, showing differences in isoform-specific expression patterns during development and in adult tissues. We delineate an essential role of rapgef1b in regulating spindle pole integrity and architecture coupled with faithful chromosome segregation during early mitoses. Further, our results highlight that rapgef1b is indispensable for embryo survival, maintaining progenitor fate in cranial neural crest cells and head size. Remarkably, rapgef1b also aids in orchestrating presomitic mesoderm patterning and somitogenesis via modulation of Wnt/ β-catenin signaling. This study positions rapgef1b as a pivotal regulator of early embryogenesis, integrating β-catenin -mediated lineage specification of paraxial mesoderm and its derivatives.

Results

Rapgef1 is expressed during embryonic development and in adult tissues of zebrafish

Zebrafish paralogs, rapgef1a, and rapgef1b, located on chromosomes 8 and 21 respectively, show high homology in the coding regions and encode proteins with similar domain organization (Fig. 1A). Both paralogues contain the highly conserved Crk binding region (CBR) which includes a hotspot for additional exons (orange arrowhead, Fig. 1A). The expression of rapgef1 during embryonic development and in adult tissues was examined using primers corresponding to the sequence in the 5’ end (N-ter), for the amplification of rapgef1a and 1b specific gene products (Fig. 1B, S1A). These primers are common to multiple isoforms and are predicted to amplify products of 130 bp and 167 bp from rapgef1a and 1b, respectively. Semi-quantitative PCR resulted in amplicons of expected sizes from all the developmental stages (32-cell to 2 days post fertilization, dpf). Our results show that transcripts from both paralogs are expressed during early developmental stages, such as the 32-cell stage, indicating that rapgef1 mRNA is maternally deposited. The expression of rapgef1 continues into the mid-blastula stage (256 cell- 3.3hpf stage) and somitogenesis phase (6–21somite stage) until 2dpf (Fig. 1B). A recent report has shown that depletion of Rapgef1a results in gross developmental defects, brain and vasculature abnormalities [13]. However, the role of other paralog, Rapgef1b remains unknown. Hence, we focused on investigating the developmental functions of this paralog Rapgef1b in early embryogenesis. First, to delineate the spatiotemporal and developmental gene expression profile of rapgef1b, we used in situ hybridization (ISH) with sequence-specific antisense riboprobes, which revealed its ubiquitous expression across developmental stages (red asterisk, Fig. 1C), with higher anterior expression at 1dpf (red arrowhead, Fig. 1C). Specificity of the rapgef1b expression was confirmed by using sense riboprobes as control (Fig. 1C). Western blotting showed the expression of a 150kD polypeptide of Rapgef1, similar to that seen in mammalian cells and tissues, in all the early developmental stages (Fig. 1D). Brain tissue from adult zebrafish expressed a distinct polypeptide, indicating tissue-specific expression of alternate isoforms as described earlier using mouse tissues and alluding to isoform specific role in neural functions (Fig. S1B) [18].

Fig. 1.

Expression profile of rapgef1 and its isoforms during embryogenesis and adult zebrafish tissues. A: Schematic illustration showing the protein domains of zRAPGEF1a and zRAPGEF1b, triangle (orange) indicates hotspot for additional exons in the CBR region. Numbers indicate total amino acids and the position of insertion of additional amino acids in the alternate isoforms. B: Schematic representation of rapgef1a and 1b 5’ end with the primer positions (black arrows) and the exon numbers. RT-PCR analysis of rapgef1a and 1b transcripts in different developmental stages. Zebrafish rpl7 (ribosomal protein L 7) was the loading control. NTC represents no template control. C: Whole-mount RNA in situ hybridization showing rapgef1b expression during early development. Scale bars, 200 μm. D: Western blot showing Rapgef1 levels during embryonic development. β-Tubulin and Gapdh were used as the loading controls. E: Schematic representation of rapgef1a and b exons showing predicted alternatively spliced transcripts for rapgef1a and 1b with nucleotide ID (https://www.ncbi.nlm.nih.gov/nuccore). Locations of isoform-specific primers (black arrows) are shown that give products of different lengths based on the presence or absence of additional exons in alternately spliced isoforms. F: RT-PCR analysis of rapgef1 transcripts in different developmental stages using rapgef1a and 1b isoform-specific primers. β actin was used as the loading control. G: Isoform-specific amplification of rapgef1a and 1b transcripts in adult tissues (brain, heart, liver, ovary, fin, muscle). Zebrafish rpl7 was used as the loading control. NTC represents no template control

Differential isoform-specific expression of rapgef1 paralogs in embryonic and adult tissues

Expression of a higher molecular weight isoform of Rapgef1 in the adult zebrafish brain (Fig. S1B) indicates generation of alternate isoforms, including a set of cassette exons, may be expressed. Zebrafish rapgef1 transcripts span 3257 bp (rapgef1a), and 3566 bp (rapgef1b), with the longest variants are composed of 25 (rapgef1a) and 28 (rapgef1b) exons. Publicly available databases like NCBI suggest the generation of alternatively spliced isoforms through the inclusion of a pair of exons (10 & 11 in rapgef1a and 12 & 13 in rapgef1b). Isoform-specific primers for both rapgef1a and 1b which are expected to generate PCR products of different sizes based on the inclusion of one or both exons, were used to study the expression of alternate isoforms (Fig. 1E). Rapgef1a transcript lacking the additional exons (X4, 405 bp product) is expressed throughout embryonic development till early somitogenesis (6 somite stage) (Fig. 1F). Interestingly, expression of the longer isoforms of rapgef1a with 1 or more exons, (X3-498 bp, X2-570 bp, X1-633 bp amplicons) begins with late somitogenesis (18–21 somite stage) (Fig. 1F). The longest isoform with exons 10 & 11 is predominantly expressed in the juvenile head and adult brain, with poor expression of other products suggesting the isoform-specific function of rapgef1a in the adult brain (Fig. S1C). Other adult tissues, such as the ovary, heart, liver, and kidney show expression of various isoforms in addition to the embryonic isoform (Fig. 1G).

In contrast, only the smallest isoform lacking exons 12 and 13 (X5, 200 bp amplicon) of rapgef1b is expressed until 2dpf and its levels decrease as the embryo develops (Fig. 1F). Remarkably, there is dynamic isoform switching which results in the expression of the brain-specific, longer isoform of rapgef1b containing exons 12 & 13 in the juvenile and adult brain with age (Fig. S1C). Adult tissues such as the heart, liver, ovary, and kidney predominantly express the isoform lacking cassette exons, with no detectable levels of the longer transcripts of rapgef1b (Fig. 1G). We also investigated the expression patterns of the brain-specific isoforms by ISH with sequence-specific antisense riboprobe (Fig. S1D). Rapgef1b (antisense riboprobe to exon 3 and 4) is expressed in the diencephalon (double red asterisk), rods, and cones (red arrow), ganglion cell layer (red arrowhead), gut, and myotomes of 3dpf larvae (red asterisk) (Fig. S1D). In contrast, the brain-specific isoform expression is restricted to diencephalon, and the ganglion cell layer of the eye, but is absent in rods and cones and myotome (red arrow, Fig.S1D). Thus, the expression of the brain-specific isoform of rapgef1b is restricted to the cranial region and absent from mesodermal-derived tissues like the myotome.

Rapgef1b depletion results in neurodevelopmental and anteroposterior axis patterning defects

To determine the functional role of rapgef1b in embryogenesis, we used two loss of function strategies, the knockdown approach using sequence-specific antisense morpholino (ATG/ translational blocker) and a CRISPR-based knockout approach against zRapgef1b. The morpholino approach was used for rapid and effective knockdown of gene expression to understand developmental functions during embryogenesis. Further, as morpholino-based knockdown strategy may induce off target effects, we used the CRISPR-based gene editing method and generated crispants (F0) to corroborate the morphant phenotypes (Fig. 2). The rapgef1b morphants injected with ATG blocker morpholino and rapgef1b crispants showed depletion of Rapgef1 protein. The rapgef1b KO fish showed a deletion of 366 bp, confirming effective abrogation of its expression (Fig. 2C, S2A). Both the strategies resulted in similar phenotypes, the morphants were characterized as P0 (normal, similar to WT), P1 (brain and axis defects), and P2 (severe brain and axis deformities) (Fig. 2B and D, S2D). Rapgef1b depletion by both strategies resulted in defects in the brain, eye, and inner ear in 24-hour post-fertilization embryos. The P2 embryos appear smaller in size, showed poorly developed eyes, defects in the brain architecture, and lack a proper midbrain-hindbrain boundary (red asterisk, Fig. 2B). The crispants and morphants also showed somitic defects (red arrowhead, Fig. 2B) and anterior-posterior body axis abnormalities, marked by shortening of the axis and tail (red arrow, Fig. 2B). We have used time matched, severe phenotype embryos classified as P2 to perform the comparative gene expression analyses in this study. The severity of the phenotype and the survival rate were observed in a morpholino concentration-dependent manner with time (Fig. S2B, S2C). To ascertain that the phenotype was specific to Rapgef1b knockdown, we performed rescue experiments by introducing human rapgef1 mRNA, which resulted in partial, but significant rescue (39%) of the severe phenotype, P2 (Fig. 2D, E). In contrast, human rapgef1 mRNA of a predicted deleterious mutant Y485C (Mutagenetix & ClinVar), a residue conserved in zRapgef1, that shows altered stability and compromised phosphorylation by Abl (unpublished findings from our lab), could not rescue the rapgef1b-specific phenotype (Fig. 2D, E). These results suggest that rapgef1b is essential for early development and morphogenesis of the brain, eye, somites, as well as the elongation of the body axis and tail.

Fig. 2.

Rapgef1b is required for brain and anteroposterior axis development. A: Schematic representation of rapgef1b knockdown and knockout strategies by morpholino and CRISPR-Cas9-based approaches respectively. The morpholino binds to the translation start site, abrogating protein synthesis. sgRNA was designed to generate a deletion of 366 bp between exons 3 and 4. B: Representative images showing gross morphological appearance of rapgef1b depleted embryos by morpholino and CRISPR strategies in comparison to control embryos at 1dpf showing midbrain-hindbrain boundary (red asterisk) and body axis defects (red arrow). The boxed areas show the magnified images of defects in midbrain-hindbrain defects (red asterisk) and body axis defects (red arrow). Scale bars, 500 μm. C: Upper panel showing Rapgef1 levels by western blot in rapgef1b morpholino injected morphants and rapgef1b crispants compared to WT control. β-Tubulin was used as the loading control. The lower panel shows PCR-based analysis to validate the exonic deletion in CRISPR mutants. D: Quantification showing percent phenotype of morphants in rescue experiment. Data are shown as mean ± SEM, n = 3 biological repeat experiments, with a minimum of 60 embryos analyzed for each experiment. E: Rescue experiments showing the functional equivalence of human Rapgef1 as a rapgef1b ortholog in zebrafish embryos

Rapgef1b is critical for the maintenance of progenitor fate in cranial neural crest, formation of cranial neural crest derivatives, and maintenance of head size

As rapgef1b is expressed in early development and its depletion resulted in brain and anteroposterior axis abnormalities, we inferred that rapgef1b is required for early neurodevelopment and axis specification. To test this hypothesis, we performed gene expression analysis of key cranial neural crest specification markers and patterning on rapgef1b morphants and crispants. The morphants/crispants showed a significant reduction in in the expression of the neural crest specifier, twist1a (red asterisk, Fig. 3A) and the neural plate marker sox2 (red arrowhead, Fig. 3D). Twist1a is expressed in the cranial neural crest-derived head mesenchyme, diencephalon, and is required for the migration of neural crest cells and their differentiation [29]. Sox2 is required for otic and epibranchial placode induction, promotes the maintenance of pluripotency, and regulates the development of the sensory epithelium and neural tube [30, 31]. The reduced expression of twist1a and sox2 strongly indicates that rapgef1b plays a key role in the maintenance of progenitor population in the cranial neural crest, sensory placodes, and is involved in neural tube patterning. We also observed a reduced expression of premigratory neural crest marker, sox10 (red arrow, Fig. 3B) [32, 33]. The expression was rescued in the morphants injected with human Rapgef1 mRNA (red arrowhead, Fig. 3B). These results suggest that pre-migratory neural crest cells marked by sox10 expression are reduced in the morphants and may be the underlying cause of a poorly developed brain (Fig. 2B). We also observed increased cell death in the anterior region of the embryo (forebrain, midbrain-hindbrain boundary/ MHB, and midbrain) as shown by acridine orange staining (white asterisk, Fig. S3A).

Fig. 3.

Rapgef1b is essential for cranial neural crest specification and maintenance of head size. Gene expression analysis by in situ hybridization in control and rapgef1b morphants/ mutants at 1dpf. A: Twist1a expression (red asterisk) in control and rapgef1b morphants/ mutants. B: Expression of sox10 in rapgef1b morphants, mutants (red arrow), and rescue embryos (red arrowhead). Scale bars, 200 μm. C: Pax2a expression in rapgef1b morphants/ mutants showing expression in the optic vesicle, midbrain-hindbrain boundary (red asterisk, inset), and otic vesicle (inset). D: Expression of sox2 in the neural tube of rapgef1b morphants (red arrow). Scale bars, 200 μm. E: Control and rapgef1b morphants showing post-mitotic neuronal marker neuN (white asterisk). Scale bars, 200μm. F: Skeletal preparations of rapgef1b morphants (red asterisk, inset) indicating bone and cartilage staining using Alizarin red S and Alcian blue respectively. Scale bars, 500μm. G: Quantification of microcephaly features – Head area (HA), Interpupillary distance (IPD), and Iris distance (ID) in rapgef1b morphants. All data are shown as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. n = 3 biological repeat experiments and the sample size is indicated in each image

Other key regulators of specification, differentiation, and morphogenesis are the Pax family members. Pax2a is expressed in the eye, MHB, otic placode, and the pronephros [34–37]. Pax2a, along with pax8 is required to maintain otic cell fate, and high levels of pax2a promote otic differentiation [38]. The rapgef1b morphants exhibit an expansion of pax2a expression domains in the eye, otic vesicle, and MHB (red asterisk, Fig. 3C). Thus, rapgef1b is essential to maintain progenitor cell fate, and its depletion results in an increase in differentiation-inducing signals such as pax2a. Concomitantly, we also observed an upregulation of egr2b, which is a rhombomere 3 and 5 specifier in the segmented hindbrain (Fig. S3B) [39–42]. We also examined whether rapgef1b is essential for neuronal differentiation. Neuronal nuclei (NeuN) mark the post-mitotic neurons as it is distributed in the nuclei of mature CNS neurons. Thus, NeuN marks terminal differentiation and determination of neuronal phenotype [43, 44]. The morphants showed a decrease in NeuN-positive cells (white asterisk, Fig. 3E), indicating that depletion of rapgef1b results in the decrease of post-mitotic mature neurons and strongly indicates its requirement in terminal neuronal maturation and differentiation.

At larval stages of development (4-5dpf), the rapgef1b morphants exhibited cardiac edema, severe craniofacial skeletal anomalies and defects in ceratobranchial skeletal elements (red asterisk, Fig. 3F). Craniofacial skeletal elements are derived from the sox10-positive cranial neural crest, and the mesenchyme is derived from sclerotome and lateral plate mesoderm [45]. The rapgef1b-depleted embryos showed a reduction in sox10-positive neural crest cells, which may be the underlying basis of the observed craniofacial defects. Further, the 5dpf larvae also showed microcephaly-associated craniofacial features such as reduced overall head area (HA), interpupillary distance (IPD), and iris distance (ID) (Fig. 3G), suggesting that rapgef1b plays a key role in formation of craniofacial skeleton and maintenance of overall head size in the embryo.

Rapgef1b plays an important role in paraxial mesoderm differentiation

To further explore the developmental functions of rapgef1b, we performed transcriptome sequencing analysis to identify differentially expressed genes (DEGs) and altered processes and pathways in the rapgef1b morphants contributing to the observed phenotypes. The principal component analysis (PCA) and correlation matrix showed variance and the agreement of the three replicates across conditions (Fig. S4A, B). The volcano map showed the distribution of DEGs − 76 genes were downregulated and 106 genes were upregulated at the transcriptional level with clear patterns of clustering in the rapgef1b deficient embryos within the FDR adjusted p-value < = 0.05 (Fig. 4A). The altered DEGs are listed, and their P values are shown in Fig. S4C. We also performed GO pathway analysis to further investigate the functional relationships between the DEGs. Gene ontology analysis showed an enrichment of anterior/posterior pattern specification, somite development, segmentation, and mesodermal morphogenesis genes indicating that rapgef1b plays a key role in paraxial mesoderm specification, patterning and differentiation (Fig. 4B). The heat map shows the selected up-regulated and down-regulated genes in these mentioned processes (Fig. 4C). To validate the transcriptome analysis, we assayed transcript levels of some of the up and down-regulated genes in control and rapgef1b morphants (Fig. 4D-H). The rapgef1b morphants showed increased transcript levels of the up-regulated group of genes –msgn1, tbx16, cdx4, tbx6 and bbc3. Msgn1 and tbx16 induce the differentiation program in the presomitic mesoderm (PSM) cells by reducing the expression of progenitor maintenance markers fgf8 and ntl, and inducing the expression of tbx6. Hence, msgn1 is essential to regulate the progenitor pool population until somitogenesis is complete [46, 47]. Rapgef1b morphants showed upregulation of msgn1 and tbx16 (Fig. 4D, red asterisk, Fig. 4E). We observed an overall increase in tbx6 transcripts in the morphants, particularly in the tailbud region (red arrow, Fig. 4E). Tbx6 plays important role in the specification of paraxial mesoderm, somite patterning, and myogenesis [48–50]. Studies show that ectopic tbx6 expression promotes less mesodermal fate [51]. This increase in tbx6 expression levels results in reduced mesodermal fate and may be the underlying cause of the failure of somite segmentation in the tailbud mesoderm. Thus, upregulation of msgn1 and tbx16 in rapgef1b morphants by transcriptomic analysis shows that progenitor fate is suppressed and an untimely differentiation program is initiated upon rapgef1b depletion (Fig. 4D and E).

Fig. 4.

Comparative transcriptome and differential gene expression analysis indicating the role of rapgef1b in paraxial mesoderm differentiation. A: Volcano plot of the Differentially expressed genes (DEGs) in control and rapgef1b morphants. Red dots represent DEGs with fold change (> 1 or < -1) with p-value < 0.05. Green dots represent genes that are significant with fold change (> 1 or < -1) with a p-value > 0.05. Blue dots represent genes with insignificant fold changes with p-value < 0.05. Black dots represent the genes that show insignificant levels of DEGs expression by standards of p-value and log fold change. B: Dot plot representing the Gene ontology (GO) analysis of genes upregulated in the rapgef1b morphants. C: Heatmap of Differentially expressed genes (DEGs) in control and rapgef1b morphants. The colors are mapped to z-score normalized values of the log fold change across each gene set. D: Validation by qPCR of msgn1, tbx16, cdx4 and tbx6 upon rapgef1b depletion. Data are shown as mean ± SEM, *p < 0.05, **p < 0.01, ***p < 0.001. n = 3 for each biological repeat experiment. E: Gene expression patterns of cdx4 (red asterisk), tbx6 (red arrow), msgna1 (red asterisk) and tbx16 (red asterisk) in rapgef1b morphants compared to control. Scale bars, 200 μm. F: Validation by qPCR of bbc3 upon rapgef1b depletion. G: Validation by qPCR of myhc4, myoz1b, and pavlb1 upon rapgef1b depletion. H: Validation by qPCR of her13 on rapgef1b depletion. All data are shown as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. n = 3 for each experiment. Each n represents 100 embryos

Early mesodermal cell fate is also regulated by the cdx family, which also plays key roles in axial elongation, anteroposterior patterning, and the development of the trunk and tail [52]. Differentiation of paraxial mesoderm and somitogenesis requires other tbx family members as well, along with Wnt and FGF signalling [48, 51, 53–55]. Cdx4 is also an early trunk neural crest specifier, expressed in the trunk neural crest cell progenitors and the developing tailbud [56, 57]. Interestingly, rapgef1b-depleted embryos show an upregulation of cdx4 transcript levels in transcriptome analysis, qPCR and in situ hybridization (Fig. 4C and D, red asterisk, Fig. 4E), suggesting that trunk neural crest cells are maintained in the progenitor fate in the morphants. This led us to conclude that rapgef1b is essential to initiate differentiation fate in the trunk neural crest population. Our results show that rapgef1b is essential to maintain the progenitor cell fate of the cranial neural crest (Fig. 3), however, it is required for the initiation of the differentiation fate in the mesodermal lineage (Fig. 4).

We also observed upregulation of intrinsic apoptotic genes such as bbc3 in our transcriptome analysis and qPCR validation (Fig. 4F) [58, 59]. Hence, pro-apoptotic mRNAs are upregulated in the rapgef1b morphants, which results in increased apoptotic cells, also marked by acridine orange in the cranial region of rapgef1b morphants (Fig. S3A). In contrast, rapgef1b morphants showed reduced transcript levels of skeletal muscle genes such as myhc4, myoz1b, and pvalb1 (Fig. 4G) [60–62]. Further, her13 which is required for the formation of all somitic borders was also downregulated in the morphants (Fig. 4H). The differences in expression of the above genes is likely the underlying basis of disrupted somite architecture in the rapgef1b morphants (red arrow, Fig. 2B) [63]. Hence, transcriptome analysis reveals that rapgef1b plays a crucial role in the development of paraxial (somitic) mesoderm and possibly, its derivatives.

Rapgef1b is required for presomitic mesoderm patterning and somitogenesis

Transcriptome analysis revealed that rapgef1b plays an important role in somitogenesis and mesoderm development. Hence, we examined somite development and the myogenic program in rapgef1b morphants at 1dpf. The phalloidin 568 staining showed disruption of the F-actin positive muscle fibres in the rapgef1b morphants (Fig. 5A) [64]. To ascertain the role of rapgef1b in myogenesis and somite patterning, we examined the morphants for myoD1 expression. MyoD1 is expressed in the somites and is required for the establishment and maintenance of somite-derived muscle progenitor lineages [65, 66]. Rapgef1b morphants showed defects in the somite organization and architecture. The somites appeared enlarged and irregular with imperfect boundaries (red arrowhead, Fig. 5B) as compared to the periodic and sharp somite boundaries in control (red dotted line, Fig. 5B). These somitic defects were rescued in the morphants injected with hRapgef1 mRNA (Fig. 5B). We also examined the patterning of somitic mesoderm. Tbxta is expressed in the mesoderm during early development and neuromesodermal progenitors that give rise to the spinal cord and paraxial mesoderm of the trunk and tail. At later stages, tbxta expression is restricted to the notochord and tailbud [51, 56, 67–69]. The 1dpf morphants showed notochordal defects and expanded expression of tbxta in the tailbud (red arrowhead, Fig. 5C). The diffused and disrupted somitic boundaries were also observed at the early somitogenic phase (red arrowhead, Fig. 5D) coupled with the loss of segmental myoD1 expression (red asterisk, Fig. 5E). We also observed reduced expression of myoD1 in the PSM at the bud stage, indicating that rapgef1b depletion resulted in impaired presomitic mesoderm patterning (red arrow, Fig. 5E). Further, we observed expanded tbxta expression at the bud and 6 somite stage as well (red arrow, Fig. 5F) which suggests that neuromesodermal progenitor pool is maintained in the morphants and rapgef1b is required to induce mesodermal differentiation.

Fig. 5.

Rapgef1b depletion results in defective somite formation and deregulation of genes involved in presomitic mesoderm specification. A: Phalloidin staining in 1dpf control and morphant embryos. Inset shows the organization of F actin muscle fibers. B: myoD1 expression at 1dpf showing somite architecture (red arrowhead, red dotted line in inset) in morphants and rescue embryos Scale bars, 200 μm. C: Tbxta expression in the notochord and tailbud (red arrowhead) of rapegf1b morphants at 1dpf. Scale bars, 200μm. D: Bright field images of control and rapgef1b morphants at the 6 somite stage (red arrowhead). E: myoD1 expression in control and morphants at 6 somite stage (red asterisk) and bud stage (red arrow). F: tbxta gene expression at bud stage (red arrow), and somite stage (red arrow). Scale bars, 200 μm. G: Expression of her1 in control and rapgef1b morphants in anterior somites (red asterisk) at the 6 somite stage and tail PSM (red arrow, red asterisk) at the 18 somite stage. H: Expression of her7 in early somitogenesis in control and rapgef1b morphants showing reduced expression in anterior somites (red arrow). I: Papc expression pattern in control and morphants showing reduced expression in anterior somites (red arrowhead, red asterisk), PSM (red arrow) at the 6 somite stage, and PSM (red arrowhead) at the 18 somite stage. J: Fgf8 expression pattern in control and morphants showing midbrain-hindbrain boundary (red asterisk) at the bud stage and in the head (double red asterisk), somites (red arrowhead), and tailbud (red arrow) at 14 somite stage

Somites are derived from the segmentation of the PSM under the regulation of the oscillatory gene network, which involves her genes, Wnt, notch, and FGF signalling [63, 70]. Her1 and her7 are expressed as two stripes in the PSM, tailbud, and are essential for somite formation [71]. Her deficient embryos show enlarged somites with weak boundaries. Both these genes have partial functional redundancy, yet her1 is important for the formation of anterior somites and her7 for posterior somites [53, 72, 73]. The rapgef1b morphants show a lack of her1 expression in the anterior somites in a two-stripe manner as compared to control (red asterisk, Fig. 5G). At later stages, the embryos also showed reduced expression in the tail region (red arrow, Fig. 5G). We also observed reduced expression of her7 in the somites and PSM (red arrow, Fig. 5H). Thus, the loss of segmental stripes of her1 and her7 suggests that the segmentation clock is disrupted in the PSM of rapgef1b morphants. Papc expressed in dorsal mesodermal cells during gastrulation contributes to the convergence movements and PSM [74–77]. The anterior somites (red asterisk, red arrowhead, Fig. 5I) and PSM expression of papc is absent in the morphants (red arrow, Fig. 5I). Further, papc positive cells in the paraxial mesoderm are not detected in the tailbud of morphants at somite stage (red arrowhead, Fig. 5I). This downregulation of papc in rapgef1b depleted embryos suggests impaired convergence movements in the mesoderm, ultimately resulting in somitic defects. Another PSM marker, fgf8 plays a key role in PSM maturation and somite formation [78]. Rapgef1b morphants show an expansion of fgf8 expression in the midbrain-hindbrain boundary at the bud stage (red asterisk, Fig. 5J). At later stages, posterior somites appear enlarged with imperfect somitic boundaries (red arrowhead) and expanded expression in the head (double red asterisk) and tailbud (red arrow, Fig. 5J). Hence, rapgef1b is essential for PSM patterning, expression of segmentation clock genes and somite architecture.

Rapgef1b regulates mesoderm specification by modulating canonical Wnt signaling

To investigate the underlying cause of observed paraxial mesodermal defects upregulation, we examined the Wnt/β-catenin signaling pathway in the rapgef1b morphants. Wnt signals are required for mesoderm induction, somite formation, and patterning through the β-catenin-Lef1/TCF pathway [48, 53, 79]. The abrogation of canonical Wnt signaling impairs somitogenesis and presomitic segmentation [48, 50]. Indeed, during mid-blastula stage, we observed a decrease in the Wnt signaling effector β-catenin in the nucleus and cortex in interphase and mitotic phase (white asterisk, Fig. 6A). Western blotting showed a reduction in phospho GSK-3β levels (ser9) indicative of enhanced kinase activity, in turn resulting in increased phospho β catenin levels (ser33/37) in rapgef1b morphants (Fig. 6B). Consequently, total β catenin levels decreaed in the morphants (Fig. 6B), indicating the downregulation of canonical Wnt signaling. In addition, the rapgef1b-depleted embryos showed decreased protein levels of the Wnt effector axin2 (Fig. 6B). The transcript levels of Wnt effectors axin2, lef1 and cyclin D1 were also reduced in the morphants, suggesting an overall downregulation of the canonical Wnt responses (Fig. S3F). Previous studies show that RAPGEF1 localizes to the centrosome and regulates proper centriole duplication in mammalian cells in vitro [5]. Hence, we examined the role of rapgef1b in centrosomal dynamics during embryonic mitoses, at the mid-blastula stage. The morphants exhibited severe centrosomal and centriolar defects as shown by γ-tubulin (white asterisk, Fig. 6C, S3E) and centrin 3 respectively, (Fig. S3C). Hence, rapgef1b is required to maintain spindle pole integrity, centrosome structure and organization during early embryonic mitoses (Fig. 6C). The rapgef1b-depleted embryos also showed elongated spindle length (red dotted line, Fig. 6C) and chromosome congression anomalies at metaphase (Fig. 6C). These defects are likely to result in impaired mitotic progression and chromosome instability in rapgef1b morphants. This is also indicated by an overall decrease in plk1 transcript levels (Fig. S3D), essential for mitotic progression, faithful segregation of chromosomes, and proper spindle assembly [80]. Interestingly, we observed a partial rescue of the pericentriolar defects as shown by γ-tubulin in the rapgef1b morphants injected with human β-catenin mRNA (white asterisk, Fig. 6D). These results demonstrate that the centrosomal functions of rapgef1b, particularly the maintenance of spindle pole integrity are mediated by Wnt β-catenin signaling.

Fig. 6.

Rapgef1b functions by modulating Wnt/β catenin signaling during early embryonic development. A: Sum projection of confocal images of 256 cell stage control and rapgef1b morphants embryos showing β-catenin (white asterisk) during interphase (upper panel) and mitosis (lower panel). Scale bars, 50 μm. n = 3 for each experiment. Each n represents 50 embryos B. Western blotting analysis showing Rapgef1, Gsk3β/phospho Gsk3β levels, and β-catenin/phospho β-catenin levels in control and rapgef1b morphants. β-Tubulin and Gapdh were used as the loading controls. C: Sum projection confocal images of 256 cell stage control and rapgef1b morphants embryos showing centrosomal staining by γ tubulin (green) and DNA (blue/DAPI). Quantification of spindle length (red dotted line) and chromosome congression defects in the rapgef1b morphants as compared to control. Scale bars, 50 μm. All data are shown as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. n = 3 for each experiment. Each n represents 40 embryos. D: Sum projection confocal images of 256 cell stage embryos injected with rapgef1b morpholino and β-catenin mRNA showing centrosomal staining (white asterisk) by γ tubulin (green) and DNA (blue/DAPI). β-catenin mRNA injected embryos serve as control. Scale bars, 50 μm. Quantification of spindle pole integrity (white asterisk) in the rapgef1b morphants as compared to control. All data are shown as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001. n = 3 for each experiment. Each n represents 40 embryos. E: Quantification showing percent phenotype of morphants when treated with Wnt activator and Wnt inhibitor. F: Gene expression patterns of tbx6 (red asterisk, red arrowhead) in control, rapgef1b morphants, morphants treated with Wnt inhibitor and activator. WT embryos treated with Wnt activator only and inhibitor only were used as positive control. G: Bright field images of control and rapgef1b morphants at the 18 somite stage (red asterisk). H: Gene expression patterns of myoD1 (red asterisk, red arrowhead) in control, rapgef1b morphants, morphants treated with Wnt inhibitor and activator. WT embryos treated with Wnt activator only and inhibitor only were treated as positive control. All data are shown as mean ± SEM, n = 3 for each biological repeat experiment. Each n represents 20 embryos

To further probe rapgef1b-mediated Wnt signaling in mesoderm patterning, we exposed morpholino-injected embryos to the Wnt activator CHIR99021 or the Wnt inhibitor XAV939 (Fig. 6E). Morphants exposed to XAV939 resulted in an increased number of P1 and P2 phenotypes (red arrow, Fig. 6E) which were rescued by administration of CHIR99021, akin to the introduction of human rapgef1 mRNA (blue arrow, Fig. 6E). Further, expression of the paraxial mesoderm specifier tbx6 was also restored in rapgef1b-depleted embryos injected with rapgef1 mRNA (red asterisk, Fig. 6F) and CHIR99021 (double red asterisk, Fig. 6F). Conversely, rapgef1b depleted embryos treated with XAV939 showed an additive phenotypic expression of tbx6 (red arrow, Fig. 6F). The somitic defects were also observed in rapgef1b morphants at the 18 somite stage, marking the later phase of somitogenesis (red asterisk, Fig. 6G, H). The severity of the somitic defects increased upon treatment with the Wnt inhibitor, reinforcing the synergistic action of rapgef1 loss and attenuated Wnt signaling (double red asterisk, Fig. 6H). The somite organization and architecture were restored upon treatment with Wnt activator (red arrow, Fig. 6H). Further, inhibition of Wnt signaling also results in reduced axis elongation [81], which is consistent with the reduced Wnt signaling in rapgef1b morphants. To summarise, rapgef1b depletion reduces canonical Wnt/β-catenin signaling, which is the underlying basis for the impaired mesodermal patterning and somitic defects.

Discussion

Embryogenesis is characterized by dynamic spatiotemporal gene expression that drives progenitor fate specification and subsequent differentiation. As Rapgef1 regulates cell adhesion, cytoskeletal organization, and cell proliferation, it is well poised to play key roles in embryonic development. However, its precise role in embryogenesis remains unclear, largely because murine Rapgef1 null embryos fail to progress beyond implantation. Hence, we investigated rapgef1 functions in zebrafish embryos, which have two paralogs rapgef1a and rapgef1b. Rapgef1a depletion resulted in reduced survival rate, gross morphological defects in vasculogenesis, CNS development, and reduced locomotor activity [13]. These zebrafish paralogs may have unique as well as overlapping functions; however, the contribution of rapgef1b in biological processes remains unknown.

We show that both paralogs undergo tissue- and stage-specific alternative splicing, with isoform switching evident across developmental transitions. In contrast to rapgef1a, the short isoform of rapgef1b is exclusively expressed across developmental stages, strongly indicating its unique, non-overlapping functions during early development (Fig. 1E). Further, the short rapgef1b isoform predominates during proliferative phases of brain development, whereas longer isoforms containing cassette exons are enriched in the adult brain, where differentiation is complete. Longer isoforms with cassette exons show differences in intra-molecular interactions which have been predicted to alter GEF activity [19]. This temporal and tissue-specific isoform expression suggests a mechanism for fine-tuning Rapgef1-mediated signaling and GTPase regulation during embryogenesis. The evolutionary conservation of splicing hotspots across vertebrates further supports the functional differences between the isoforms. This also highlights some of the limitations of our current study, as isoform-specific reagents such as antibodies are not available. Future studies employing isoform-specific reagents will be critical to dissect the unique and redundant roles of these transcripts in embryonic development and to highlight their evolutionary significance. Differences in temporal expression of isoforms generated from the two paralogs may be due to the presence of alternate regulatory elements on the two linkage groups, and suggests that their products play independent roles.

Our study provides the first evidence that rapgef1b is essential for lineage specification in the cranial neural crest and paraxial mesoderm (Fig. 7). Using CRISPR-Cas9-mediated knockout and morpholino knockdown approaches, we demonstrate that loss of rapgef1b disrupted early embryogenesis, neurodevelopmental and anteroposterior axis elongation defects (Fig. 2). Rapgef1b crispants and morphants showed reduced expression of neural crest specifier genes twist1a and sox10, increased apoptosis, and resulted in smaller head size. Twist1a and sox10 mark the undifferentiated cranial neural crest and regulate the specification, maintenance of the progenitor pool, and migration [32, 82–84]. Further, loss of sox10 results in apoptosis and failure to undergo differentiation to form CNS and cranial neural crest-derived craniofacial skeleton [33, 82, 85]. In addition to the specification of neural crest cells, rapgef1b also plays a key role in the formation of neural crest derivatives. We observed the absence of neural crest-derived melanocytes in morphants and crispants (Figs. 2 and 3). The transcriptomic profile also showed differential expression of genes involved in melanocyte differentiation and pigmentation (Fig. S4D). Rapgef1b morphants showed severe craniofacial abnormalities that may be attributed to cardiac edema and the reduced expression of sox10, failing to undergo differentiation to form craniofacial bones and cartilage. Depletion of rapgef1b also resulted in mitotic abnormalities, including defective spindle pole integrity and chromosome mis-congression in zebrafish blastulae. Centriole amplification has been shown to affect the proliferation and survival of neural progenitor cells in zebrafish [86]. These results reveal that the centrosomal functions of rapgef1 are also conserved during embryonic mitoses in zebrafish, in a manner akin to that in mouse and human cells [5]. Together, the reduced neural crest gene expression in rapgef1b morphants, coupled with enhanced apoptosis may be a consequence of impaired mitosis. The small head size, shortened interpupillary distance, mitotic aberrations, and increased apoptosis allude to a microcephaly-like phenotype in rapgef1b morphants. Together, these findings suggest that Rapgef1b is required to sustain progenitor identity in cranial neural crest cells and maintain mitotic fidelity during early development. Further, it is imperative to delineate the mitotic regulation by rapgef1b in the embryonic brain to gain mechanistic insights into its neurodevelopmental functions.

Fig. 7.

rapgef1b regulates neural crest maintenance, mitotic fidelity, and paraxial mesodermal differentiation during zebrafish embryogenesis. Schematic summary of rapgef1b functions in early development. On the left (blue), rapgef1b maintains neural crest progenitors and supports neural tube and Midbrain–hindbrain boundary (MHB) patterning (twist1a, sox10, pax2a, sox2), which contribute to craniofacial skeletogenesis and neuronal differentiation (sox2, NeuN). In the centre (orange), rapgef1b ensures mitotic fidelity and spindle pole integrity (cen3, plk1), enabling cell survival and proper head size maintenance (sox10, bbc3a). On the right (green), rapgef1b modulates canonical Wnt/β-catenin signaling to regulate Presomitic mesoderm (PSM) patterning (msgn1, tbx16, her1, her7, papc, tbxta, fgf8), somitogenesis (tbx6, myoD1, her13, fgf8), and myogenic differentiation (myhc4, myoz1b, pvalb3). Together, these findings establish rapgef1b as a key regulator linking progenitor maintenance, mitotic control, and mesodermal tissue differentiation

Beyond the CNS, our RNA-seq and expression analyses demonstrates that rapgef1b also governs paraxial mesoderm patterning, somitogenesis, and axis formation. Similar to Drosophila Rapgef1, which is required for larval muscle integrity, zebrafish rapgef1b morphants exhibited notochordal and somitic defects [26, 87]. These defects correlated with reduced expression of presomitic mesoderm (PSM) genes, particularly in anterior somites, consistent with disrupted segmentation. The PSM is derived from trunk mesodermal progenitors, and these progenitor cells are maintained in an undifferentiated state by activation of Wnt signaling and tbxta expression [88, 89]. Canonical Wnt signals are key regulators of PSM maturation and somite formation by regulating the expression of lineage determinants [78]. Mammalian RAPGEF1 interacts with β-catenin as well as GSK3β, modulating Wnt signaling [7, 8]. The rapgef1b depletion results in the downregulation of Wnt signals, disrupting the balance between proliferation and differentiation, which induces premature pro-differentiation signals in mesodermal progenitor cells. Hence, rapgef1b morphants show disrupted PSM specification and defects in somite organization and architecture. Given that PSM progenitors are maintained by canonical Wnt signaling, our pharmacological experiments show that rapgef1b modulates this pathway, potentially via GSK3β and β-catenin regulation. Rapgef1b depletion dampened Wnt activity and decreased expression of Wnt effectors, leading to premature differentiation signals in mesodermal progenitors. In addition, decreased Wnt/β-catenin signaling reduced the expression of neural crest specifiers, showing that Wnt signaling coupled with BMP signals is essential for the expansion of neural crest progenitors and maintains their multipotency [90]. The downregulation of Wnt/β-catenin signaling also resulted in a surge of apoptosis in the CNS, consistent with prior reports linking Wnt loss to cell death in the midbrain and cerebellum [51, 91–94]. In fact, β-catenin signaling promotes differentiation of neural crest cells into pigment cell types and suppresses neuronal cell fates in premigratory neural crest. Intriguingly, the transcriptome profiling of rapgef1b morphants also showed downregulation of genes involved in melanocyte differentiation and pigmentation, along with an overall decrease in neural crest-derived melanocytes in morphants and crispants. This is suggestive of rapgef1b-mediated attenuation of β-catenin signaling, resulting in decreased expression of melanocyte differentiation-associated genes.

In summary, our study involving detailed spatiotemporal gene expression analyses, transcriptome profiling and pharmacological intervention experiments establish rapgef1b as a critical regulator of cranial neural crest and mesodermal lineage specification, acting in part through modulation of Wnt signaling. The data highlight the role of rapgef1b in maintaining progenitor states, ensuring mitotic integrity and survival during early development. Our findings show that rapgef1b paralog is essential for early development, though there is the presence of a duplicate gene with high homology. Yet, we do observe some overlapping redundant phenotypic attributes such as reduced survival rate, gross morphological brain and body axis defects in rapgef1b crispants, as reported for rapgef1a morphants [13]. Thus, a detailed analysis of the unique and redundant functions of the two paralogs of rapgef1 in zebrafish is imperative to gain mechanistic insights into the developmental significance of rapgef1 vasculogenesis, somitogenesis and motor neuron activity. Besides the independent functions of the two paralogs, the phenotypic effects shown by the knockdown or knockout approaches may be due to the insufficient dosage effects. Lastly, as lineage specification is also regulated by chromatin remodeling, a function attributed to RAPGEF1 in mammalian cells, it will be interesting to determine how rapgef1b regulates chromatin modifications to alter the expression of lineage determinants during zebrafish embryogenesis.

Methods

Zebrafish lines, MO injection, and characterization of phenotypes

Tubingen strain (TU-AB) zebrafish were raised according to standard protocols as described earlier. All experiments were performed according to protocols approved by the Institutional Animal Ethics Committee of the Council of Scientific and Industrial Research, Centre for Cellular and Molecular Biology, India. All the experiments were performed for a minimum of three biological replicates, and statistical analyses are stated in the figure legends. Embryos were obtained from the natural spawning of adult fish, kept at 28.5°C, and staged according to hours after fertilization [95]. The endogenous rapgef1 levels were depleted by using morpholino (rapgef1b translation blocker, 5’-CTTGCTTTCTATTTTCCCCGACATG-3′ (Gene Tools), 0.25mM, 0.5mM and 1mM in each embryo and standard control MO, 5′-CCT​CTT​ACC​TCA​GTT​ACA​ATT​TAT​A-3′ (Gene Tools), 0.25mM, 0.5mM and 1mM was used as negative controls. All the assays were performed with 1mM translation blocker unless stated otherwise. For rescue experiments, 50pg of human RAPGEF1 or β-CATENIN mRNA was co-injected along with rapgef1b MO in each embryo at the one-cell stage. The embryos were then analyzed for gross morphological defects and survival at later stages of development (1dpf).

Generation of the rapgef1b knockout line

Single guide RNAs (sgRNAs) targeting rapgef1b (Ensembl ID: ENSDART00000123759.3) for CRISPR/Cas9-mediated knock-out were designed using CRISPOR online tool (www.https://crispor.tefor.net/). Two sgRNAs targeting the exons 3 and 4 of the rapgef1b gene were used with the following target sequences: 5′-GAGATACTTTAAGACGATTG − 3′ and 5′-GCTTGGCGACTCTGATTCGC-3′. Gene-specific oligos were annealed in a PCR machine, ramping temperature down from 95 °C to 4 °C in a stepwise manner and run on agarose gel electrophoresis to ensure correct annealing. Both the sgRNAs were cloned into pDR274 plasmid (Addgene #42250). RNAs from both clones were in vitro transcribed using the Maxiscript SP6/T7 kit (AM1322) according to the manufacturer’s instructions. One-cell stage zebrafish embryos were injected with 3nl of a solution containing 600pg of Cas9 protein (TrueCut™ Cas9 Protein v2 #A36498) and 150pg of each sgRNA. The CRISPR/Cas9 approach generated a deletion of 366 bp, resulting in a premature STOP codon in exon 4. For genotyping, genomic DNA was obtained by incubating the samples (whole embryos or adult caudal fin fragments) in TE buffer supplemented with 1mM EDTA and 10 µg/ml Proteinase K (Cat no. 193981) for 1 h (embryos) or 4 h (fins) at 55 °C and 10 min at 95 °C, and then stored at 4 °C. Primers sequences used for genotyping can be found as Fig S5 [96].

RNA isolation, semi‑quantitative RT‑PCR, and sequencing

Total RNA was isolated from 100 embryos for each group using the RNA isolation kit (MN; 740955.50) as per the manufacturer’s protocol. Total RNA from various zebrafish adult tissues was prepared using RNAiso Plus (TaKaRa, cat. no. 9109) according to the manufacturer’s protocol. cDNA was synthesized from total RNA using PrimeScript 1^st strand cDNA synthesis kit (TaKaRa, cat. no 6110 A). PCR was carried out using the prepared cDNA for amplification of zebrafish rapgef1 isoforms and rpl7 (internal control). Zebrafish isoform-specific primers for rapgef1a and rapgef1b were used to identify splice variants of these isoforms present in embryonic and adult zebrafish tissues. All primers used in this study are listed in Fig. S2. PCR was performed in BioRad C1000 Touch thermal cycler with, Taq DNA Polymerase (TaKaRa, cat. no. R500A). The following PCR conditions were used for amplification of rapgef1: initial denaturation was at 94 °C for 5 min, followed by 30 cycles of denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s, and extension at 72 °C for 30 s. The final extension was for 10 min at 72 °C. Amplified PCR products were examined and pictures were captured using the Vilber-Lourmat Gel Documentation system (Germany). The PCR fragments were eluted using Macherey-Nagel NucleoSpin Gel and PCR clean-up kit (MN; 740609.50) as per the manufacturer’s protocol and sequenced to confirm that the amplicons are generated from alternate splice variants. All primers used in this study are listed in Fig.S5.

Antisense riboprobe preparation

For cloning sequence-specific exonic fragments to synthesize riboprobes, total RNA was isolated from 1dpf embryos of the TU-AB strain using an RNA isolation kit (MN; 740955.50). The total RNA was used to prepare cDNA using the PrimeScript 1^st strand cDNA synthesis kit (TaKaRa, Cat. # 6110 A). Sequence-specific primers were used to amplify the specific gene fragment and cloned into the pGEMT easy vector and the sequence was verified. The sequence-verified plasmid was linearised using Nco1 and antisense digoxigenin (DIG)-labeled riboprobes were synthesized using the DIG RNA labeling kit (Roche #11175025910) [97]. All primers used in this study are listed in Fig.S5.

Whole mount RNA in situ hybridization

Embryos of various stages were fixed in 4% paraformaldehyde overnight followed by methanol fixation. Methanol-fixed embryos were rehydrated with 50% MeOH/PBT (1× PBS and 0.1% Tween 20). Embryos were rinsed with PBT, a 1:1 PBT/hybridization wash, and then a hybridization wash (50% formamide, 1.3 × SSC, 5 mM EDTA, 0.2%, Tween-20, H2O). Embryos were incubated with a hybridization mix (50% formamide, 1.3 × SSC, 5 mM EDTA, 0.2%, Tween-20, 50 µg/mL yeast t RNA, 100 µg/mL heparin, H2O) for 1 h at 55 °C. The riboprobes prepared in the hybridization mix were added and incubated for 16 h at 55 °C. Then, embryos were rinsed with prewarmed hybridization wash for 30 min (2 times) at 55 °C and followed by 1 × TBST wash (5 M NaCl, 1 M KCl, 1 M Tris, pH 7.5, and 10% Tween 20) at room temperature. Then, blocking was done with 10% FBS heat-inactivated fetal bovine serum (Gibco; 16210-064) for 1 h at room temperature. The embryos were then incubated with an Anti-DIG-AP antibody (Roche 11093274910) with a dilution of 1:5000 overnight at 4 °C. The next day, embryos were transferred into a 6-well plate and washed twice with 1X NTMT (0.1 M NaCl, 0.1 M Tris-Cl at pH 9.5, 0.05 M MgCl2, 1% Tween-20 and H2O) for 10 min; then, incubated with 1:1 NBT-BCIP solution (Cat no. PI34042) NTMT solution for the color development. The embryos were monitored during this color reaction and stopped with 1× PBT as a stop solution. Images of embryos were captured using a Zeiss Stemi 2000CVR bright-field microscope at 5× magnification (with AxiocamICc1) [98] The embryos after in situ hybridization at 3dpf were cryoprotected with 30% sucrose in PBS, embedded in Tissue freezing medium (Leica) and cryosectioned at 15 μm thickness.

Western blot analysis

Whole zebrafish embryos (n = 100) and adult tissues were homogenized using RIPA buffer (20mM Tris–HCl, pH 7.4; 150mM NaCl; 5mM EDTA; 1% NP-40; 0.4% sodium deoxycholate, 0.1% SDS and 1× Protease inhibitor cocktail) and kept on ice for 15 min with vortexing 3 times. The homogenized lysates were centrifuged (12000 rpm for 20 min at 4 °C), and supernatant were taken for the analysis. Subsequently, samples were boiled in 1× Laemmli buffer and subjected to SDS-PAGE. 20 µg of total protein was loaded in each lane on 8% SDS-PAGE gel for detection of endogenous proteins. Proteins were transferred to PVDF membranes (Merck Milipore, ISEQ00010) by electro-blotting. Membranes were blocked for 2 h at room temperature in TBST (20 mM Tris HCl, pH 7.5, 150 mM NaCl, 0.1% TWEEN 20) containing 3% BSA and probed overnight with primary antibodies (Fig. S6). After three washes with TBST, the blots were incubated with HRP-conjugated secondary antibody for 1 h, and then washed in TBST. The blot was developed for chemiluminescence signal using the chemiluminescence (Bio-Rad) and captured in the Vilber-Lourmat Chemiluminescence System (Germany). Densitometry analysis was done using Image-J. β-Tubulin was used as a control for the normalization of protein in all the samples. The data were plotted as mean normalized protein level relative to WT ± SD was plotted as error bar. Data from at least three independent experiments were plotted and significance was calculated with unpaired t-test using Prism 8.0. All the antibodies used are listed in Fig. S6.

Real-time PCR

For quantitative real-time PCR (RT-PCR), we performed PCR using Power SYBR Green PCR Master Mix (Applied Biosystems) with ViiA 7 Real-Time PCR System (Applied Biosystems) according to the manufacturer’s instructions. Quantification data are represented as means ± SEM of three independent experiments. Analyses on normalized data were performed using the 2 − δδCT algorithm (the delta−delta-Ct or ddCt algorithm). All genes were normalized against zrpl7 unless mentioned otherwise. The GraphPad Prism software was used for the analysis. For the statistical significance of the data, p-values were calculated by performing an unpaired Student t-test. All primers used in this study are listed in Fig.S5.

Immunohistochemistry of zebrafish embryos

Embryos of various stages were fixed in 4% paraformaldehyde overnight followed by methanol fixation. Methanol fixed embryos were rehydrated with 50% MeOH/PBST (1X PBS and 0.1% Triton X-100) followed by incubation in blocking solution 1% BSA/PBST for 2 h at room temperature. Then the embryos were incubated with primary antibody overnight at 4 °C. Next day 1X PBST washes, appropriate secondary antibody incubation overnight at 4 °C. After washing, DAPI (1 µg/ml) was added for 10 min followed by washing with 1X PBS and stored at -20 °C until confocal/fluorescence imaging [98]. The mitotic phenotypes were quantified using 3D reconstruction of the confocal images using the IMARIS software. All the antibodies used are listed in Fig. S6.

Acridine orange staining

Acridine orange (AO) staining was performed to detect the apoptotic cells in 1dpf embryos. Embryos were dechorionated, and stained with 5 mg/L AO for 1 h in the dark followed by washing with embryo water. Then the embryos were anesthetized with tricaine and were photographed by a Zeiss AxioZoom.V16 microscope, the bright green dots in the embryos indicate apoptotic cells. The number of apoptotic cells in the head region was counted using Image J software [99].

Skeletal preparations by alcian blue and alizarin red staining

Wild-type control and morphants 5-day-old larvae were fixed in 4% PFA/PBS overnight at 4 °C. The embryos were washed with 1XPBS and stored overnight in 100% methanol at -20 °C. The following staining solutions were prepared - Solution A (0.04% Alcian blue, 125mM MgCl2 in 70% ethanol) and Solution B (1.5% Alizarin Red S). 1 ml of solution A and 10ul of solution B were added to the larvae and kept overnight at room temperature with constant rocking. The staining solution was removed and the larvae were washed two times with water, followed by bleach solution for 1 h at room temperature and a gradation of Glycerol-KOH. The embryos were then stored in 50% glycerol/ 0.25% KOH for 2 h at room temperature. The stained larvae were then imaged using a Zeiss Stemi508 stereomicroscope. For microcephaly analyses of zebrafish 5dpf larval head area was defined by the otic vesicle and the semicircle of eyes as posterior and lower boundary, respectively. The interpupillary distance (IPD) and distance between irises are also measured and quantitated using ImageJ software [100].

Phalloidin staining

Embryos were dechorinated and fixed using 4% paraformaldehyde for overnight at 4 °C. Embryos were washed out of 4% paraformaldehyde at least three times with Phosphate Buffered Saline-0.1% Tween20 (PBT). Prior to phalloidin staining, embryos were permeabilized for 30 min at room temperature with PBS-1%Trition. Embryos were incubated in Phalloidin–Tetramethylrhodamine B isothiocyanate (cat no.P1951) at 1∶500 dilution overnight at 4 °C. Embryos were washed three times in PBS-0.1%Tw prior to imaging.

Pharmacological modulation of the Wnt/β-catenin signaling pathway

Zebrafish embryos were treated at shield stage with 10 µM XAV939 and 10 µM CHIR99021 dissolved in DMSO, to inhibit (XAV939) or activate (CHIR99021) Wnt/β-catenin signalling. At 24hpf the percent phenotype was calculated for all these treated groups of embryos.

RNAseq-based gene expression analysis

The total RNA was extracted from 1dpf control and rapgef1b depleted embryos using an RNA isolation kit (MN; 740955.50). Sequencing libraries were generated using MGIEasy RNA library prep set (MGI) according to the manufacturer’s instructions. Here, 500 ng total RNA was used as starting material, and rRNA was depleted by MGIEasy rRNA depletion Kit. The rRNA-depleted samples were fragmented, reverse transcribed and the second strand was synthesized and converted to cDNA. DNA was then purified using DNA Clean Beads provided in the kit followed by end repair and A-tailing. Barcoding and adaptor ligation were performed and the samples were purified. Samples were amplified using adaptor-specific primers and quantified using a Qubit dsDNA high-sensitivity kit (ThermoScientific). Sample fragment size was determined using 4200 TapeStation (Agilent). 1pMol of the dsDNA library was denatured and circularized to make single-stranded circular DNA. DNA Nano Balls (DNBs) were created using Rolling cycle amplification. DNBs were loaded onto patterned flowcell and sequenced using PE100 recipe on MGISEQ-2000 sequencer (MGI).

The raw sequence data from the control and rapgef1b-depleted embryos were evaluated for read quality status using FASTQC [101]. The RNAseq data was uploaded on Gene Expression Omnibus (GEO), accession no. GSE283261. Adapter trimming and read filtration were performed using CUTADAPT, and the filtration criteria were set to a minimum length (50 bp), and a minimal quality score (Q20) cutoffs. The trimmed reads were then mapped onto the zebrafish fish reference genome (GRCz11) using STAR aligner [101]. STAR was run in quant-mode, instructing the tool to generate the read counts and alignments. The differential gene expression was performed using the DESeq2 package [102], assayed across the three replicates of each condition; control (WT) and rapgef1b knockdown (rapgef1b KD). Transcripts with a raw count sum of less than 10 were excluded for downstream analysis. A correlation matrix was calculated to determine the agreement of various replicates across conditions. To ensure consistency across conditions and the replicates, principal component analysis (PCA) was carried out of deseq2 normalized counts. Significantly differentially expressed transcripts were obtained by filtering for a false discovery rate (FDR) by filtering p-adjusted cutoff of p < = 0.05. These are then classified as upregulated and downregulated based on their respective log fold change (L.F.C) values, which is representative of their regulation states; transcripts that had a fold change greater than 1 (L.F.C > = 1) were considered as up-regulated while those lesser than − 1 (F.C<=-1) were deemed down-regulated. Gene ontology (GO) enrichment analysis was performed using the enrichGO module of the ClusterProfiler package to analyze the effect of these gene subsets on metabolic networks [103].

Authors’ contributions

TP performed the morpholino and CRISPR experiments, PCR, ISH, and analyzed the data, prepared the figures, and assisted in manuscript writing. SI performed the cloning of ISH markers, ISH, immunohistochemistry, imaging, and analyzed the data. SD performed ISH, imaging, and analyzed data. RJM analyzed the transcriptome data and prepared the figure. DTS supervised the transcriptome analysis. VR and MK developed the concept. TP, VR, and MK designed the experiments and interpreted the data. MK supervised and mentored the entire study and wrote the manuscript with input from other authors.

Funding

TP and SI acknowledge CSIR, India for the research fellowship. VR thanks CSIR for the award of ES scheme, 21(1089)/19/EMR-II. MK thanks the Council of Scientific and Industrial Research- Centre for Cellular and Molecular Biology (CSIR-CCMB), Govt. of India, and the Department of Science and Technology (DST), Govt. of India (DST/INSPIRE/04/2016/001436) and the Anusandhan National Research Foundation (ANRF), Govt. of India (ANRF/ECRG/2024/000070/LS) for support and funding this research.

Data availability

No datasets were generated or analysed during the current study.

Declarations

Competing interests

The authors declare no competing interests.