Splicing is a crucial regulatory node of gene expression that has been leveraged to expand the proteome from a limited number of genes. Indeed, the vast increase in intron number that accompanied vertebrate emergence might have aided the evolution of developmental and organismal complexity. Here, we review how animal models for core spliceosome components have provided insights into the role of splicing in vertebrate development, with a specific focus on neuronal, neural crest and skeletal development. To this end, we also discuss relevant spliceosomopathies, which are developmental disorders linked to mutations in spliceosome subunits. Finally, we discuss potential mechanisms that could underlie the tissue-specific phenotypes often observed upon spliceosome inhibition and identify gaps in our knowledge that, we hope, will inspire further research.

One of the major evolutionary transitions between prokaryotes and eukaryotes was the emergence of introns, which resulted in the fragmentation of previously contiguous protein-coding genes. The presence of introns required the co-evolution of a splicing machinery, called the spliceosome, of which there are two types: major and minor. Each core machinery consists of five small nuclear RNAs and over 150 associated proteins, many of which are shared between the major and minor spliceosomes (Table S1). These splicing machineries act analogously to splice major and minor introns, respectively.

The maintenance of introns and spliceosomes has energetic costs, which potentially explains the general trend towards intron loss across evolution (Rogozin et al., 2012). One exception to this trend is the vertebrate lineage, which possess a relatively high number of introns (Rogozin et al., 2012). This suggests that splicing is uniquely leveraged by vertebrates as a regulatory mechanism. Indeed, splicing can be used to create temporal delays in protein production, to titrate protein levels and to expand the proteome from a limited number of genes through alternative splicing (AS). Thus, splicing might have played a major role in generating the complexity observed in vertebrates.

In this Review, we focus on the role of splicing in vertebrate development, with an emphasis on its role in key processes such as cell cycle regulation, cell fate specification and cell differentiation. We also present our current understanding of the role of splicing in organogenesis, as evidenced in animal studies. The tissue-specific requirement for spliceosome components is further revealed by spliceosomopathies, which are disorders linked to pathogenic variants in spliceosome components. Therefore, we also briefly discuss developmental spliceosomopathies and the potential mechanisms that might underlie their tissue-restricted symptoms.

Cell cycle regulation

Given the central role of splicing in gene expression, it is no surprise that cell cycle progression relies on spliceosome function. Screens for genes that are essential for cell division have especially underscored the importance of splicing for mitotic progression (Martín et al., 2021; Mu et al., 2014; Neumann et al., 2010; Sundaramoorthy et al., 2014). Inhibition of ∼30 core spliceosome components has been shown to result in prometaphase delay followed by misalignment of chromosomes in metaphase. These defects might stem, at least in part, from intron retention in CDCA5, the gene encoding sororin, which is essential for sister chromatid separation. As a consequence of this retained intron, CDCA5 transcripts are degraded by nonsense-mediated decay (NMD), resulting in the reduction of sororin protein levels (van der Lelij et al., 2014; Watrin et al., 2014). Indeed, ectopic expression of full-length, properly spliced sororin transcripts in cells deficient for components of the spliceosomal NineTeen Complex (NTC), is able to rescue mitotic arrest (Watrin et al., 2014).

The importance of splicing in cell cycle progression is further illustrated by the finding that several key cell cycle genes are regulated through splicing changes that occur in a cell cycle phase-dependent manner. RNAseq of synchronized HeLa cells revealed that 15% of all expressed genes are subject to periodic cell cycle-dependent AS (Dominguez et al., 2016). This includes aurora kinase B (AURKB), a mitotic checkpoint protein that is crucial for the binding of chromosomes to kinetochores and for chromosomal segregation. In G2 phase, a high percentage of AURKB mRNA transcripts contain a retained intron, resulting in their degradation by NMD. In this way, AURKB protein levels are regulated such that they peak in M phase (Dominguez et al., 2016). Although this example illustrates the significance of splicing in cell cycle regulation, the functional importance of many other cell cycle phase-dependent AS events remains unexplored.

Cell cycle phase-dependent splice pattern shifts suggest differential spliceosome activity. This is likely achieved through post-translational modifications (Fig. 1A), as spliceosome components are not differentially expressed throughout the cell cycle (Santos et al., 2015). Indeed, several spliceosome components are phosphorylated by cyclin-dependent kinases (CDKs), the activity of which oscillates throughout the cell cycle (Morgan, 2007). For example, CDK1-mediated phosphorylation of SF3B1 peaks at G2/M and decreases as the cell progresses through G1/S (Murthy et al., 2018). Similarly, the NTC component cell division cycle 5-like (CDC5L) is phosphorylated by CDK2 in a cell cycle-dependent manner (Gräub et al., 2008). This phosphorylation is not only essential for splicing, but also for efficient G2/M transition (Gräub et al., 2008). Finally, phosphorylation of AS factors such as SRSF10 can also control cell cycle progression (Fig. 1B). At steady state, the majority of SRSF10 is phosphorylated and acts as a sequence-specific splicing activator (Feng et al., 2008). However, upon dephosphorylation, its function changes and it acts as a splicing repressor by inhibiting formation of the spliceosome A complex (Shin and Manley, 2002). This dephosphorylation of SRSF10 occurs in M phase and is responsible for the general splicing repression that takes place in mitosis (Shin and Manley, 2002). Moreover, it has been shown that SRSF10 becomes dephosphorylated upon DNA damage, resulting in a splicing switch from the pro-survival to the pro-apoptotic isoform of Bclx (BCL2L1) (Shkreta et al., 2016). Together, these data put SRSF10 at the center of DNA damage checkpoints during G1/S and G2/M transitions, and illustrate how splicing can dictate whether a cell is fated to live or die.

Fig. 1.

Splicing in cell cycle regulation and lineage specification. (A) Spliceosome activity can be regulated during the cell cycle via the phosphorylation of spliceosome components by cyclin-dependent kinases (CDKs). (B) Phosphorylation of the splicing factor SRSF10 regulates alternative splicing of the pro-apoptotic gene Bclx. (C) The role of alternative splicing in cell differentiation and lineage specification. Examples are shown for the genes FOXP1, BIN1 and PAX6.

Fig. 1.

Splicing in cell cycle regulation and lineage specification. (A) Spliceosome activity can be regulated during the cell cycle via the phosphorylation of spliceosome components by cyclin-dependent kinases (CDKs). (B) Phosphorylation of the splicing factor SRSF10 regulates alternative splicing of the pro-apoptotic gene Bclx. (C) The role of alternative splicing in cell differentiation and lineage specification. Examples are shown for the genes FOXP1, BIN1 and PAX6.

Lineage specification and differentiation

One of the first pieces of evidence that suggested that AS could play a role in regulating cell fate emerged following the identification of mutually exclusive exons in the gene encoding the FOXP1 transcription factor (Fig. 1C) (Gabut et al., 2011; Han et al., 2013). These exons are alternatively spliced upon the differentiation of human embryonic stem cells (hESCs): a pluripotent cell type derived from the epiblast. The FOXP1 isoform expressed in ESCs is required for the expression of pluripotency genes, such as NANOG and OCT4, and represses the expression of differentiation genes, thereby revealing the potential for AS to regulate pluripotency (Gabut et al., 2011). More recently, transcriptomic analyses of hESCs and differentiated cell types of endodermal, mesodermal and ectodermal origin revealed that ∼20% of expressed genes undergo AS during hESC differentiation (Fagg et al., 2022; Xu et al., 2018). Many of these AS events are distinct between endoderm-, mesoderm- and ectoderm-derived cell types, suggesting that splicing plays a crucial role in producing these lineages (Fig. 1C). Indeed, siRNA-mediated knockdown of core spliceosome components in ESCs represses pluripotency markers and often activates the expression of totipotent markers (Rodriguez-Terrones et al., 2018; Shen et al., 2021; Strikoudis et al., 2016). Similarly, treatment with pladienolide B, which inhibits the spliceosome by binding to the SF3B complex, leads to intron retention in pluripotency genes but transcriptional activation of totipotency genes (Shen et al., 2021). Together, these findings suggest that splicing plays a crucial role in restricting potency, thereby facilitating lineage diversification and the production of organismal complexity.

The mechanism by which the spliceosome regulates cell potency remains unclear. However, it has been noted that key totipotent genes have fewer and shorter introns than pluripotent genes (Shen et al., 2021), whereas genes expressed in terminally differentiated cells, such as voltage-gated ion channels, contain even more introns. As such, splicing might be especially important for the expression of genes associated with a restricted cell fate. This is further underscored by the observation that knockdown of spliceosome components in mouse and human pluripotent stem cells specifically results in AS and downregulation of genes involved in neuronal differentiation (Laaref et al., 2020; Shen et al., 2021). Interestingly, embryoid bodies deficient in the spliceosome component Htatsf1 fail to differentiate into neuroectoderm, while mesoderm and endoderm form efficiently (Corsini et al., 2018). Thus, the spliceosome might be particularly important for regulating a neuroectoderm fate.

Insights from constitutive knockout studies

Given that many spliceosome components are essential for cell cycle progression, cell survival and cell differentiation in cell culture, it is not surprising that constitutive knockout of several spliceosomal components results in early embryonic lethality in mice (Fig. 2A; Table S2). Although many spliceosome components are essential for pre-implantation development in mouse, zebrafish lacking spliceosome components survive at least up to 2 days post-fertilization (dpf), when gastrulation has already completed (Fig. 2B; Table S2). For example, even though knockout of the minor spliceosome component Rnpc3 in mice results in death before blastocyst formation around E3.5, mutation in rnpc3 in zebrafish results in lethality between 7 and 10 dpf, which is well into organogenesis (Doggett et al., 2018; Markmiller et al., 2014). This variability in the timing of lethality might be explained by differences in the scale and timing of the maternal-to-zygotic transition in these species. For example, it is known that early embryonic development is predominantly controlled by maternally deposited mRNAs and proteins, which in mice and humans are degraded by the 4-cell stage, but exist for longer in zebrafish (Vastenhouw et al., 2019). Accordingly, low levels of fully spliced, maternally deposited rnpc3 mRNAs are detected up to 24 h post fertilization (hpf) in rnpc3-mutant zebrafish (Markmiller et al., 2014). Consequently, these larvae experience a delayed effect of rnpc3 loss-of-function on splicing until after the gastrulation stage. Overall, these animal studies underscore the importance of spliceosome components in pre-implantation development. However, as knockouts for only ∼20% of spliceosome subunits have been produced so far (Table S2), it remains unclear whether all spliceosome components are essential for early embryonic development. Additionally, it is important to recognize that early embryonic lethality precludes further studies into the role of splicing in vertebrate development. Therefore, the generation of conditional knockout (cKO) models, whereby a splicing factor of interest is depleted in a spatially and/or temporally restricted manner, will be important to further our understanding of splicing in development (see Box 1 for more information on the role of AS in development).

Box 1. Myotonic dystrophy: the first example of AS regulating development

One of the first pieces of evidence for the role of AS in development comes from research focused on understanding the molecular defects underpinning myotonic dystrophy. Myotonic dystrophy is an autosomal dominant disease that is caused by (CTG)n expansions in the 3′ untranslated region of DMPK (Brook et al., 1992; Mahadevan et al., 1992). The number of expansions increases with each generation, correlating with disease severity (Wong et al., 1995). It has been shown that DMPK transcripts containing (CUG)n expansions form nuclear foci in which AS factors known as muscleblind proteins (MBNL1, MBNL2 and MBNL3) are sequestered (Davis et al., 1997; Miller et al., 2000). Thus, myotonic dystrophy is caused by the indirect loss of MBNL function (Kanadia et al., 2003). These MBNL proteins regulate the AS of genes essential for skeletal muscle development by modulating the transition from fetal to adult isoforms (Lin et al., 2006). In essence, myotonic dystrophy is the result of a failed developmental transition in AS patterns. MBNL1 and MBNL2 are now known to be negative regulators of many cassette exons. For example, inhibition of MBNL proteins in differentiated cells affects splicing of the crucial pluripotency transcription factor FOXP1, and reverses them back to ESC-like state (Han et al., 2013). The crucial role of other AS factors, such as PTBP, NOVA1/2, CELF1-6 and TDP43, in development has since been elucidated and is discussed in many excellent reviews (Baralle and Giudice, 2017; Conboy, 2017; Forman et al., 2021; Liu et al., 2018; Su et al., 2018; Martí-Gómez et al., 2022).

Fig. 2.

Knockout of core spliceosome components results in early embryonic lethality. (A,B) Stages of early embryonic development with timeline for mouse (A) and zebrafish (B). Knockout of spliceosome components listed in black results in lethality at the time point indicated on the timeline. For spliceosome components listed in gray, the exact time point of embryonic lethality was not established. In zebrafish, lethality is not observed until around 2 dpf. E, embryonic day; hpf, hours post-fertilization. More details can be found in Table S2.

Fig. 2.

Knockout of core spliceosome components results in early embryonic lethality. (A,B) Stages of early embryonic development with timeline for mouse (A) and zebrafish (B). Knockout of spliceosome components listed in black results in lethality at the time point indicated on the timeline. For spliceosome components listed in gray, the exact time point of embryonic lethality was not established. In zebrafish, lethality is not observed until around 2 dpf. E, embryonic day; hpf, hours post-fertilization. More details can be found in Table S2.

The expression of major and minor spliceosome components during early development

The majority of spliceosome components are ubiquitously expressed throughout development, yet certain expression biases exist towards both specific tissue types and developmental time points. For example, genes encoding major and minor spliceosome proteins are expressed at high levels at E10.5 and E11.5 in mouse tissues and are downregulated during later developmental stages (Cardoso-Moreira et al., 2019). Moreover, RNAseq has revealed that most spliceosomal genes are enriched in the embryonic cerebrum and liver compared with other tissues, suggesting a crucial role for spliceosomes in central nervous system development and hematopoiesis (Cardoso-Moreira et al., 2019). As development occurs in three dimensions across time, explorations of the spatiotemporal expression of spliceosome components in development has frequently been investigated through whole-mount in situ hybridization. This has revealed that minor spliceosome snRNAs are enriched in the developing mouse forebrain, branchial arches, limb buds and somites (Baumgartner et al., 2015). Similarly, spliceosome components such as Eftud2 and Sf3b4 are enriched in the developing head, brain and branchial arches at E8.5, before being expressed more widely (Beauchamp et al., 2019; Yamada et al., 2020). Interestingly, no signal is observed for these components in the heart during early organogenesis (Beauchamp et al., 2019; Ruiz-Lozano et al., 1997), suggesting that this tissue might be less affected by mutations in spliceosome components.

Whole-mount in situ hybridization in zebrafish has revealed that most spliceosome components are widely expressed at early embryonic stages (An and Henion, 2012; Lei et al., 2017; Li et al., 2021; Liu et al., 2015; Markmiller et al., 2014; McElderry et al., 2019; Ríos et al., 2011; Wang et al., 2018; Yu et al., 2019). However, from 24-48 hpf, the expression of eftud2, dhx15, usp39, sf3b1, prpf31, bcas2 and prpf4 is enriched in the brain and eye, while rnpc3 expression is restricted to the eye and digestive organs. Similar observations have been made in Xenopus, where snrpb is ubiquitously expressed at the neurula stage, while eftud2, pqbp1, wbp11, sf3b4 and txnl4a are enriched at the anterior neural plate and neural crest regions (Devotta et al., 2016; Iwasaki and Thomsen, 2014; Park et al., 2022). At the tailbud stage, the expression of these spliceosome components is also elevated in the pharyngeal arches. Thus, ubiquitously expressed spliceosome components are in fact frequently enriched in neuronal and/or neural crest cells.

Given the widespread expression of spliceosome components throughout vertebrate embryonic development, it is not surprising that pathogenic variants in spliceosome subunits result in systemic developmental diseases, which are collectively known as spliceosomopathies. To date, around 50 spliceosomopathies have been reported, ranging from cancers to rare developmental disorders, such as microcephalic osteodysplastic primordial dwarfism type I (MOPD1), and neurodegenerative diseases, such as retinitis pigmentosa (Table S3). Several of these spliceosomopathies are associated with pathogenic variants in the same gene. For example, MOPD1, Roifman syndrome and Lowry-wood syndrome have all been linked to loss-of-function mutations in RNU4ATAC and can be characterized by many overlapping symptoms (Edery et al., 2011; Farach et al., 2018; He et al., 2011; Merico et al., 2015). This raises the possibility that they are not distinct disorders, but rather part of a spectrum with variable degrees of symptom severity. These complex genotype-phenotype relationships are yet to be fully understood and would benefit from biochemical studies that focus on the effects of pathogenic variants on splicing (see Box 2 for the effects of splice site mutations on disease and the potential for therapeutics).

Box 2. Splice site mutations and the potential for therapeutics

Historically, disease-causing variants have predominately been identified using whole exome sequencing. As this is often restricted to coding regions, pathogenic variants in non-coding introns or spliceosomal snRNAs frequently go undetected (Suwinski et al., 2019). With whole-genome sequencing becoming more affordable, more and more developmental Mendelian diseases are being linked to pathogenic variants in splice sites of introns, which result in aberrant splicing patterns. Splice site mutations linked to diseases are overrepresented in the 5′SS (Olthof et al., 2020), which are recognized through base pairing with U1 or U11 snRNA. Therefore, several gene therapies are currently being developed using engineered snRNAs. These strategies involve the use of a U1 snRNA with compensatory mutations that restore base pairing or exon-specific U1 snRNA, which binds to intronic sequences downstream of the 5′SS. Although cell culture experiments have demonstrated the efficiency with which these modified U1 snRNA molecules are able to rescue the aberrant splicing patterns resulting from pathogenic splice site variants (Balestra et al., 2019; Donegà et al., 2020; Lee et al., 2019; Scalet et al., 2019; Yamazaki et al., 2018), relatively limited data are available for the effects of modified U1 snRNAs in vivo. This is in sharp contrast to the use of antisense oligonucleotides, such as splice switching oligos, to rescue aberrant splicing patterns. Several of these have already been proven very successful in clinical trials for a variety of diseases and, in the case of spinal muscular atrophy and Duchenne muscular dystrophy, are even approved by the US Food and Drug Administration (Coutinho et al., 2019). In all, the use of RNA-based therapies for the treatment of diseases caused by splicing aberrations is rapidly evolving and holds great potential for the expansion of precision medicine.

The symptoms associated with spliceosomopathies have already been described in several recent reviews (Beauchamp et al., 2020; Griffin and Saint-Jeannet, 2020) and therefore are not discussed in detail here. Instead, we focus on how disruption of splicing impacts the development of organ systems that are frequently affected in spliceosomopathies. These include the ectoderm-derived nervous system and craniofacial skeleton, as well as the mesoderm-derived axial and appendicular skeleton (Fig. 3). Remarkably, the endoderm lineage, which produces most of the cells of the gastrointestinal, respiratory, genitourinary and endocrine systems, is not frequently associated with mutations in spliceosome components (Fig. 3). This might mean that endoderm-derived tissues possess specific features that make them less susceptible to splicing deficiencies than tissues derived from other germ layers. However, it must be noted that constitutive loss of spliceosome components, such as rnpc3 and ddx46 in mouse and zebrafish, do affect gastrointestinal development, raising the possibility that endoderm-derived phenotypes in spliceosomopathies are merely under-reported (Doggett et al., 2018; Hozumi et al., 2012; Markmiller et al., 2014).

Fig. 3.

Tissues affected in spliceosomopathies. Colouring (red to blue gradient) represents the percentage of spliceosomopathies that are characterized by defects in the different organ systems. Phenotype data for spliceosomopathies was obtained from Human Phenotype Ontology database.

Fig. 3.

Tissues affected in spliceosomopathies. Colouring (red to blue gradient) represents the percentage of spliceosomopathies that are characterized by defects in the different organ systems. Phenotype data for spliceosomopathies was obtained from Human Phenotype Ontology database.

Spliceosome inhibition affects cortical development and results in microcephaly

More than half of all spliceosomopathies are characterized by microcephaly (Table S3). Several other spliceosomopathies, such as early-onset cerebellar ataxia (RNU12) and pontocerebellar hypoplasia (CDC40 and PPIL1), are characterized by cerebellar defects (Chai et al., 2021; Elsaid et al., 2017). However, the high prevalence of primary microcephaly and intellectual disability among spliceosomopathies underscore the importance of splicing in neural development, particularly cortical development.

Cortical development is highly conserved across vertebrates and starts off with neurulation, the process by which the neural tube is formed from the neural ectoderm. The vertebrate neural tube is made up of neuroepithelial stem cells that initially undergo symmetric cell divisions to amplify the progenitor pool. Their location along the anterior-posterior (AP) axis informs which component of the central nervous system they will become. Neuroepithelial cells that give rise to the brain and spinal cord are first transformed into radial glial cells – multipotent neural progenitor cells. After a few self-renewing symmetric divisions, these cells switch to asymmetric differentiative divisions, producing one radial glial cell and a daughter cell with reduced potency (Noctor et al., 2004). In mammals, these differentiative daughter cells can either be another neural progenitor cell of reduced potency, called an intermediate progenitor cell, or a neuron (Noctor et al., 2004). In contrast, asymmetric differentiative divisions by radial glial cells in fish immediately result in the production of a neuron instead of producing intermediate progenitor cells. Finally, in mammals, the neurons produced during embryonic development use radial glial cells as a scaffold to migrate in an inside-out fashion to produce six distinct neuronal layers in the adult cortex.

To understand the molecular pathogenesis underlying microcephaly in spliceosomopathies, several animal models have been developed wherein splicing factors are either depleted constitutively or specifically in neural stem cells. In zebrafish, loss of eftud2, usp39, sart1 or bcas2 results in microcephaly-like phenotypes (Table S2) (Deml et al., 2015; Henson and Taylor, 2020; Lei et al., 2017; Ríos et al., 2011; Yu et al., 2019). Additionally, a forward genetic screen revealed that loss of several other splicing factors results in cell death in the zebrafish brain and spinal cord (Table S2) (Amsterdam et al., 2004). In eftud2-null zebrafish, neuronal genes are downregulated, while the expression of nestin, a neural progenitor marker, does not change (Lei et al., 2017). Microcephaly in these zebrafish is likely caused by neural progenitor cell death, leading to reduced neuron production (Lei et al., 2017).

Although the above zebrafish studies underscore the importance of splicing in neural development, little mechanistic insight has been gleaned, and most of our current mechanistic understanding of microcephaly in spliceosomopathies instead comes from loss-of-function mouse models for the spliceosome proteins Pqbp1, Sf3b4 and Ppil1, as well as the minor spliceosome snRNA U11 (Fig. 4; Table S2). Even though all these mouse models present with microcephaly at birth, the precipitation of this phenotype occurs through distinct cellular defects during embryonic development. For example, Nestin-Cre-mediated loss of Pqbp1, a component of the NTC that interacts with Sf3b1 and Txnl4a, results in an ∼10% increase in cell cycle length, primarily caused by a longer M phase (Ito et al., 2015). Even though a prolonged cell cycle length is frequently associated with increased neuron production, no changes in the number of asymmetric differentiative divisions are observed upon loss of Pqbp1 (Ito et al., 2015). Moreover, apoptosis is not elevated in Pqbp1 cKO mice, suggesting that microcephaly in PQBP1-related diseases is caused by reduced cell proliferation. Consistently, the neuron-specific ablation of Pqbp1 does not result in microcephaly (Ito et al., 2015). Together, these findings reveal the disproportionate requirement of a spliceosome component in progenitors versus post-mitotic cells.

Fig. 4.

The role of core spliceosome components in neuroectoderm development. Schematics showing the expression patterns of the spliceosome components Pqbp1, Rnu11 and Eftud2 in mouse embryos. Conditional knockout of these factors results in aberrant splicing and cellular defects affecting neuronal and neural crest cell development. This ultimately leads to gross phenotypes, such as microcephaly or craniofacial malformations.

Fig. 4.

The role of core spliceosome components in neuroectoderm development. Schematics showing the expression patterns of the spliceosome components Pqbp1, Rnu11 and Eftud2 in mouse embryos. Conditional knockout of these factors results in aberrant splicing and cellular defects affecting neuronal and neural crest cell development. This ultimately leads to gross phenotypes, such as microcephaly or craniofacial malformations.

Autosomal dominant inheritance of pathogenic variants in SF3B4 has been linked to Nager syndrome and Rodriguez acrofacial dysostosis (Bernier et al., 2012; Irving et al., 2016). The current model is that these mutations result in haploinsufficiency of SF3B4, thereby causing splicing defects that underpin the pathogenesis (Bernier et al., 2012; Petit et al., 2014). Surprisingly, Sf3b4+/− mice are mildly microcephalic from E14.5 onwards, even though neither SF3B4-related disease is characterized by microcephaly (Yamada et al., 2020). This discrepancy in phenotypes between patients and animal models suggests multiple possibilities: (1) the diseases do not result from haploinsufficiency but from a toxic gain of function of SF3B4 such that it spares cortical development; (2) the human brain is less sensitive than the developing mouse cortex to reductions in SF3B4 levels; or (3) the human developing brain expresses a factor that can compensate for SF3B4 loss of function. Further research is needed to separate these options. Regardless, the E13.5 Sf3b4+/− forebrain does not display significant differences in the number of apoptotic cells, but does exhibit a significant reduction in mitotic cell numbers (Yamada et al., 2020). Thus, cell cycle defects rather than cell death are the likely cause of microcephaly in these mice.

In contrast, cell death (of self-amplifying radial glial cells) does contribute to the microcephaly observed at E14 in U11 cKO embryos (Baumgartner et al., 2018). However, this apoptosis, which is caused by DNA damage accumulation and subsequent p53 upregulation, is not the only cellular defect underpinning microcephaly in these mice; the aberrant splicing of a subset of minor introns results in prolonged cell cycle duration, owing to both an extended S phase and prometaphase, as well as cytokinesis defects (Baumgartner et al., 2018; White et al., 2021). Although it is unclear whether this cell cycle lengthening drives U11-null radial glial cells to switch from symmetric proliferative divisions to asymmetric differentiative divisions, the cell cycle defects are a major contributor to the observed microcephaly, as this phenotype cannot be rescued by blocking p53-mediated apoptosis (White et al., 2021). Importantly, even though RNU11 has not been linked to any congenital diseases to date, these studies provide insight into the likely mechanisms underlying microcephaly in other minor spliceosome-related diseases, for example those linked to mutations in RNU4ATAC, RNPC3, CENATAC and CRIPT (Table S3).

Finally, homozygous knock-in of the hypomorphic mutation Ppil1A99T in mice results in a reduction in cerebellar size and severe microcephaly. These phenotypes mimic the symptoms observed in individuals with the same variant in PPIL1, leading to pontocerebellar hypoplasia with microcephaly (Chai et al., 2021). In contrast to other spliceosome-related microcephaly models, Ppil1A99T/A99T mice exhibit significantly reduced numbers of cortical neurons, most severely affecting the number of CTIP2+ layer V and VI neurons (Chai et al., 2021). This reduction is predominately due to p53-mediated apoptosis in immature neurons, as radial glial and intermediate progenitor cell populations are only slightly reduced. Moreover, upregulation of yH2AX is observed in the developing cortex of these mice, suggesting accumulation of significant DNA damage. RNAseq of E14.5 Ppil1 mutant brains revealed elevated retention of short and GC-rich introns, affecting the expression of genes involved in protein translation, RNA processing, the DNA damage response and axon development.

In summary, these animal models underscore the importance of the spliceosome in neural development, in part through regulating p53-mediated apoptosis, but especially in regulation of neural progenitor cell proliferation. These effects are mediated via the aberrant splicing of many regulatory genes, such as the cell cycle regulators Anapc4, Spc24 and Emx2 (Baumgartner et al., 2018; Chen et al., 2019; Yamada et al., 2020). Of note, although overexpressing canonically spliced transcripts of Anapc4 and Emx2 has been shown to rescue some of the microcephaly phenotypes, this is not predicted to result in full rescue, as spliceosome inhibition likely simultaneously affects the splicing of many genes crucial for cortical development (Chen et al., 2019; Yamada et al., 2020).

Spliceosome inhibition results in craniofacial defects

Besides microcephaly, other frequently observed defects in spliceosomopathies are craniofacial abnormalities affecting nasal, mandible, ear and calvarial structures, among others (Table S3). These skeletal and cartilaginous structures are predominantly derived from neural crest cells. Neural crest cells are multipotent cells that, after undergoing an epithelial-to-mesenchymal transition, migrate to different areas of the body and differentiate into several different cell types. For example, cranial neural crest-derived neural stem cells differentiate into cranial ganglia, while cranial neural crest-derived mesenchymal stem cells give rise to osteoblasts that eventually form craniofacial structures. Specifically, the cranial neural crest cells located most anteriorly give rise to osteoblasts that form the frontonasal skeleton, whereas those originating from the developing midbrain ultimately produce the zygomatic, maxillary and mandibular processes. In addition, neural crest-derived mesenchymal stem cells located in the hindbrain migrate to the pharyngeal arches and produce the bones and cartilage of the jaw, inner ear and neck.

The high frequency of craniofacial abnormalities in spliceosomopathies linked to pathogenic variants in components of the SF3B complex and U5 snRNP have been the driving force for studies focused on splicing in neural crest development. Consequently, most current animal models inhibit Eftud2, Txnl4a, Sf3b2 or Sf3b4 (Tables S2 and S3). Homozygous loss of eftud2 in zebrafish results in dysplasia of Meckel's cartilage, ceratohyal and ethmoid bones (Wu et al., 2019). Additionally, malformations of the sphenoidal sinus and notochord are observed and the otolith is absent in these mutants. eftud2+/− zebrafish exhibit a milder phenotype, with a shortened mandibular bone, resembling the mandibular hypoplasia observed in individuals with mandibulofacial dysostosis (Wu et al., 2019). Given that the affected craniofacial structures are formed by neural crest cells of the first, second and third branchial arch, these findings suggest that splicing regulated by Eftud2 plays a role in neural crest proliferation, migration and/or differentiation. Indeed, homozygous loss of Eftud2 in murine neural crest cells results in several craniofacial abnormalities, most notably hypoplasia of the frontonasal prominence (including mandible, maxillary and nose), absence of the middle and outer ear, and hypoplasia of the first and second pharyngeal arches (Beauchamp et al., 2021). These severe defects are attributed to apoptosis of neural crest cells found in the respective developing craniofacial regions. The molecular changes that underlie this increased apoptosis include elevated levels of exon skipping and upregulation of genes involved in the p53 pathway (Beauchamp et al., 2021). Importantly, inhibition of p53 significantly increases the size of the first pharyngeal arch in Eftud2−/− mouse embryos, suggesting that p53 activation and neural crest cell death is ultimately responsible for the craniofacial anomalies (Beauchamp et al., 2021).

Mutations in SF3B2 and SF3B4 have also been associated with craniofacial defects (Table S3). Antisense morpholino oligonucleotide-mediated inhibition of either sf3b2 or sf3b4 at the neurula stage in Xenopus significantly affects expression of the transcription factor genes sox10, tfap2 and snai2 (Devotta et al., 2016; Timberlake et al., 2021), which are known regulators of neural crest development. In contrast, the expression pattern of sox2, which marks neural plate progenitors, is expanded in both sf3b2 and sf3b4 morphant embryos. Inhibition of sf3b2 at the tailbud stage leads to a decrease in neural crest cells migrating into the pharyngeal arches (Timberlake et al., 2021). Finally, later injection of sf3b2 and sf3b4 antisense morpholinos results in hypoplastic and missing neural crest-derived cartilage in stage 45 tadpoles (Devotta et al., 2016; Timberlake et al., 2021). This reduction in neural crest cells is the consequence of increased cell death, while the cell cycle is generally unaffected in sf3b4 morphant embryos (Devotta et al., 2016). Overall, the lack of cell cycle defects and precipitation of neural crest cell death point to a common etiology of the craniofacial phenotypes observed in spliceosomopathies. Indeed, it has recently been shown that antisense morpholinos against snrpb, eftud2 or txnl4a also affect the formation of craniofacial cartilage in tadpoles through increased cell death and a concomitant reduction in neural crest gene expression, particularly sox10 (Park et al., 2022). The exact splicing defects that underlie this cell death remain unclear as no genome-wide data have been generated so far, and RT-PCR analysis did not reveal splicing defects in the neural crest genes sox10, tfap2 or snai2. However, the splicing of sox9, another neural crest marker that has been shown to regulate sox10 expression (An and Henion, 2012), is mildly affected by inhibition of sf3b4 (Devotta et al., 2016).

The role of splicing in craniofacial development might have evolutionary ramifications. Jaw evolution has been a crucial evolutionary adaptation that has allowed jawed vertebrates to exploit innumerable environmental niches. The exquisite breadth of jaw evolution is exemplified by the beaks of Darwin's finches, the snouts of anteaters and the diverse jaw structures of Cichlid fish in the East African Great Lakes. While transcriptional programs obviously play an important role in generating these diverse shapes, RNAseq analysis of the oral and pharyngeal jaws of Cichlid fish species also point to an important role for splicing. The number of genes that are differentially spliced between these species outnumbers the number of differentially expressed genes almost threefold, suggesting that specific patterns of AS play an important role in adaptive changes of the Cichlid jaw (Singh et al., 2017). Viewed from this lens, the high frequency of jaw phenotypes associated with spliceosomopathies might be a reflection of splicing being leveraged for jaw evolution.

Spliceosome inhibition results in skeletal defects

While skull bones are neural crest derived, the majority of the skeletal system is actually derived from the mesoderm. During gastrulation, the mesoderm is divided into four regions: (1) axial mesoderm that forms the notochord; (2) paraxial mesoderm that gives rise to the axial skeleton, dermis and all skeletal muscle; (3) lateral plate mesoderm that produces the appendicular skeleton and the cardiovascular system; and (4) intermediate mesoderm that forms the urogenital system. Development of the axial skeleton starts with a mesenchymal-to-epithelial transition of cells that make up the somites. These differentiate into dermatome, myotome and the sclerotome, which produces connective tissues of the axial skeleton, including vertebrae, ribs, tendons and ligaments. After their production, mesenchymal stem cells from the sclerotome migrate and surround the notochord and neural tube, which in turn induce chondrogenesis and ossification, ultimately resulting in the vertebral column. The appendicular skeleton is formed by mesenchymal cells from the lateral plate mesoderm that outgrow to form the limb bud. These mesenchymal cells then differentiate into chondrocytes to promote invasion of vasculature into the ossification center, after which point long bones are produced through endochondral ossification. Ultimately, the identity of the bones of the appendicular and axial skeleton depends on the molecular signals that are received by signaling centers that govern patterning along the three axes: proximal-distal (PD), dorsal-ventral (DV) and AP.

Defects in chondrogenesis and ossification frequently result in skeletal dysplasias, short stature or even complete absence of skeletal elements. The presence of these skeletal abnormalities in the majority of spliceosomopathies suggests that splicing is important for these cellular processes (Table S3). As such, the molecular and cellular etiology of these skeletal defects has gained more attention (Table S2).

The presence of severe limb defects, such as mesomelia, absent long bones and polydactyly, in Rodriguez syndrome points to an important role for SF3B4, and perhaps the entire SF3b complex, in skeletal development (Irving et al., 2016). The shortening or even absence of long bones specifically suggests a defect in endochondral ossification. Indeed, growth plates from fetuses with Rodriguez syndrome exhibit a reduced number of disorganized hypertrophic chondrocytes with altered morphology (Marques et al., 2016). RNAseq of chondrocytes from one of the fetuses with a heterozygous mutation in SF3B4 revealed downregulation of transcription factors that regulate skeletal development and many aberrant splicing patterns, but especially increased exon skipping (Marques et al., 2016). Given that haploinsufficiency for SF3B4 is thought to cause Nager syndrome and Rodriguez syndrome, heterozygous knockout of Sf3b4 in mice was predicted to recapitulate the limb defects observed in patients (Bernier et al., 2012). Surprisingly, heterozygous knockout mice for Sf3b1 and Sf3b4 display skeletal abnormalities affecting only the AP axis of the axial skeleton (Isono et al., 2005; Yamada et al., 2020). With relatively low penetrance, several vertebrae are transformed in these mice (e.g. L6->S1). Moreover, homeobox genes, which are known to instruct the positional identities of cells along the AP axis, are ectopically expressed in the paraxial mesoderm and second pharyngeal arch of Sf3b1+/− mice (Isono et al., 2005). As genome-wide effects of heterozygous loss of the SF3b complex on splicing and gene expression has not yet been investigated, further data are necessary to elucidate the molecular mechanism by which AP axis patterning is affected in these mice. The fact that Sf3b4 heterozygous mice do not show the profound skeletal defects observed in Nager syndrome and Rodriguez acrofacial dysostosis patients strengthens the argument that point mutations in SF3B4 might result in a toxic gain of function. However, the aforementioned caveats of potential mouse-to-human differences and redundancy mechanisms need to be formally tested.

Heterozygous loss of Wbp11 also results in axial skeleton defects in mice (Martin et al., 2020). These include vertebral fusions, hypoplastic transverse processes and butterfly vertebrae in the cervical region of the axial skeleton, but also rib defects and abnormalities of the sternum. Although these phenotypes underscore the importance of Wbp11 for axial skeleton formation, the underlying molecular and cellular events remain unclear. Homozygous loss of Pqbp1, another Prp19 complex component, in neural progenitors and in a subset of mesenchymal cells in the bone marrow of mouse embryos also results in short stature, brachycephaly and prognathism (Yang et al., 2020). Moreover, Pqbp1 cKO mice show a reduction in bone mass and bone mineralization, concomitant with reduced expression of key chondrocyte and osteoblast markers (Yang et al., 2020). In contrast, no change is observed in the number of osteoclasts. Finally, knockdown of Cdc5l in chondrocytes results in G2/M arrest and reduced expression of chondrocyte markers (Jokoji et al., 2021), which might reflect decreased numbers of chondrocytes, a differentiation defect or specific downregulation of only these markers. Thus, further experiments are needed to differentiate between these possibilities. However, given that Cdc5l depletion also results in upregulation of Col10a1, a marker of hypertrophic chondrocytes, this case likely reflects a differentiation defect (Jokoji et al., 2021). Interestingly, morpholinos against both wbp11 and pqbp1, injected into Xenopus embryos, result in truncation of the AP axis and reduced expression of mesodermal markers such as fgf4 and wnt8 (Iwasaki and Thomsen, 2014). Combined with the fact that Wbp11+/− mice also show defects in the kidneys, which are mesodermally derived, these data suggest that the prp19 complex is required for mesoderm proliferation and/or differentiation (Martin et al., 2020).

Insight into the importance of splicing in lateral plate mesoderm development comes from a cKO mouse in which U11 snRNA is ablated in the limb bud mesenchyme. Despite the fact that U11-null pups present with micromelia, i.e. shortening of the limbs, basic segmentation along the PD axis is largely unaffected in the cKO mice (Drake et al., 2020). Consistently, the expression profiles of key patterning genes such as Shh, Fgf8 and Hoxa11 are unaltered. Instead, disruption of minor intron splicing in these mice was found to particularly affect the expression of genes regulating cell cycle progression. Similar to loss of U11 in neural progenitor cells, U11 loss in chondro-osteoprogenitors results in a longer cell cycle, owing to prolonged S phase and a delay in the prometaphase-to-metaphase transition (Baumgartner et al., 2018; Drake et al., 2020). Together, this suggests that disrupted cell cycle progression might be a conserved mechanism by which minor spliceosome inhibition affects tissue development. This might also explain why distal chondro-osteoprogenitors undergo apoptosis, as these progenitors generally divide most rapidly in the limb bud (Boehm et al., 2010). Finally, although the specific effects of U11 loss on chondro-osteoprogenitor differentiation have not been investigated, reduced width and ossification of long bones suggests that the minor spliceosome is also important for mesoderm differentiation into chondrocytes and/or osteoblasts (Drake et al., 2020).

Despite their ubiquitous expression, loss-of-function of core spliceosomal components often results in tissue-specific developmental defects that affect the nervous and skeletal system (Fig. 3; Table S2). These findings suggest that core spliceosome components are not always essential for splicing, as cell culture experiments would suggest. Indeed, the tissue specificity of spliceosome function is further evidenced by the lack of reports describing defects in the endocrine and digestive system in spliceosomopathies, suggesting that the development of these organ systems is less susceptible to aberrant splicing. Although enriched expression of spliceosome components in the developing brain has frequently been reported, many of the tissue-restricted phenotypes observed in animal models and spliceosomopathies cannot be explained by this expression bias. Interestingly, this phenomenon is akin to that seen in ribosomopathies, which are disorders characterized by mutations in ubiquitously expressed ribosomal components. It remains unclear why some tissues are more affected than others upon splicing inhibition. Several animal models have revealed that constitutive loss of spliceosome components not only differentially affects the development of tissues, but also cell types within a tissue. For example, loss of U11 snRNA in the developing mouse forebrain predominantly affects self-amplifying radial glial cells, while those undergoing differentiative divisions or intermediate progenitor cells and neurons are seemingly less prone to inhibition of minor intron splicing (Baumgartner et al., 2018). So why are some cell types more affected by loss of spliceosome components than others? Although this is a major outstanding question in the field that requires much more research, we can speculate on some potential explanations (Fig. 5).

Fig. 5.

Potential mechanisms underlying cell type- and tissue-specific phenotypes observed upon ubiquitous spliceosome inhibition. Explanations for cell type-restricted phenotypes observed in animal models and spliceosomopathies may include cell-intrinsic differences, such as transcription and/or splicing kinetics, or differences in proliferation rates. Other causes could include differential expression of genes highly dependent on spliceosome function or differential expression of factors that can compensate for the loss of function of some spliceosome components. Finally, the differential activity of downstream pathways frequently affected by aberrant splicing, such as p53 signaling, could contribute to differences in cellular defects.

Fig. 5.

Potential mechanisms underlying cell type- and tissue-specific phenotypes observed upon ubiquitous spliceosome inhibition. Explanations for cell type-restricted phenotypes observed in animal models and spliceosomopathies may include cell-intrinsic differences, such as transcription and/or splicing kinetics, or differences in proliferation rates. Other causes could include differential expression of genes highly dependent on spliceosome function or differential expression of factors that can compensate for the loss of function of some spliceosome components. Finally, the differential activity of downstream pathways frequently affected by aberrant splicing, such as p53 signaling, could contribute to differences in cellular defects.

The contribution of cell-intrinsic differences to tissue-specific phenotypes

One reason that some cell types are more affected by loss of spliceosome components than others might be that there are cell-intrinsic differences that make some cells more susceptible to splicing inhibition (Fig. 5). For example, it was recently reported that the splicing kinetics of individual introns vary between different cell lines in culture (Bedi et al., 2021). Although such kinetics have not yet been evaluated in vivo, progenitor cells in the developing embryo with elevated splicing kinetics might be more susceptible to minor perturbations in spliceosome function. As splicing occurs co-transcriptional, differences in transcription elongation rates between cell types might also play a role. Indeed, introduction of a point mutation in RNA polymerase II that reduces transcription elongation rates, results in hundreds of AS changes in mouse neural progenitors and neurons (Maslon et al., 2019). Interestingly, neural progenitor cells with slow transcription fail to self-renew but can differentiate to immature neurons (Maslon et al., 2019). This finding highlights the importance of transcription kinetics and splicing decisions in neuronal development.

Another key difference between cell types in a developing embryo is their rate of proliferation, which in turn is linked to the number of differentiated cell types they need to produce. For example, neural precursors undergo rapid expansion to account for the billions of neurons and glia that will constitute the adult brain. As discussed above, inhibition of spliceosome components often affects proliferation through cell cycle defects, via the aberrant splicing of cell cycle regulators, and several pieces of evidence suggest that splicing inhibition particularly affects rapidly dividing cells (e.g. early embryonic lethality) (Baumgartner et al., 2018; Drake et al., 2020). The precise mechanism underlying this disproportionate effect on progenitor cells is unclear. Perhaps a slow dividing cell has enough time to repair and/or escape some of the cell cycle defects caused by the accumulating splicing defects.

The expression of redundant factors can compensate for loss of spliceosome components

Another explanation for the tissue-specific phenotypes observed in animal models and spliceosomopathies is the existence of redundancies in the splicing reaction. These might also explain why mutations in different spliceosome components can result in completely different diseases affecting distinct tissues. To illustrate, loss of minor spliceosome components, such as the snRNAs U11 and U4atac (Olthof et al., 2021), but also of shared spliceosome proteins, such as Ppil1 and Eftud2 (Beauchamp et al., 2021; Chai et al., 2021), result in elevated retention of a subset of introns, but not all. The presence of spliced transcripts in samples from individuals with spliceosomopathy can partially be explained by the fact that most spliceosomopathies are the result of haploinsufficiency or, in the case of an autosomal recessive inheritance pattern, hypomorphic mutations (Table S3). These mutations lead to a reduction in function, but not a complete loss of function. Depending on the spatial and temporal expression levels of the disease-causing gene, some tissues might therefore have sufficient functional spliceosomes left to properly splice the majority of the transcripts.

It is also possible that tissues that are relatively unaffected by splicing inhibition express factors that are able to compensate for the mutated spliceosome components (Fig. 5). This could explain why fibroblasts from MOPD1 patients with pathogenic variants in RNU4ATAC show only mild defects in minor intron splicing, while lymphoblastoid cells from the same individuals contain many aberrant minor intron-containing transcripts (Cologne et al., 2019). Similarly, knockout of Zrsr2 in murine myeloid cells leads to a modest increase in minor intron retention, whereas no effect on splicing is observed in mouse embryonic fibroblasts (Madan et al., 2022). The cell type-specific changes in splicing upon Zrsr2 KO might be, in part, the consequence of variable spatial expression of Zrsr1, an intron-less imprinted gene that has been identified as a functional paralog of Zrsr2 in both mouse and human (Kitagawa et al., 1995). Indeed, although mutation of Zrsr2 does not result in any phenotype in mice, a double Zrsr2/Zrsr1 mutant exhibits early embryonic lethality (Gómez-Redondo et al., 2020). Another possibility is in line with the idea that gene duplication often does not result in a new function, but rather results in tissue-specific expression over time. Finally, it is conceivable that AS factors, the expression of which is often tissue restricted, can compensate for the lack of certain core spliceosome components. Indeed, SR proteins have been shown to be able to substitute for U1 snRNP (Tarn and Steitz, 1994). Future studies focusing on splicing mechanisms are needed to provide more insight into redundancies that exist in the splicing reaction and that might explain the tissue-specific developmental phenotypes linked to splicing inhibition.

Restricted target expression could contribute to cell type specificity of splicing inhibition

A third explanation for the cell type-specific phenotypes is the spatially and temporally restricted expression of targets that are most dependent on specific spliceosome components for their splicing (Fig. 5). Although all vertebrate introns require the spliceosome for their removal, some introns possess stronger splice sites than others and are therefore less dependent on spliceosome proteins that stabilize snRNAs-intron base-pairing. Indeed, although SF1 is essential for cell survival, its knockdown affects the splicing of only a subset of introns (Tanackovic and Krämer, 2005). Furthermore, genes that contain many introns are likely more dependent on the spliceosome for their proper expression than genes that contain only one or two introns. Notably, genes with the highest number of introns encode calcium channels, dyneins, collagen and laminin proteins, among others (Howe et al., 2021). These proteins belong to families whose members display tissue-enriched expression patterns throughout development and function in tissue-specific processes. However, it remains hard to determine the exact contribution of tissue-specific expression of target genes to developmental phenotypes, because it is currently unclear which genes are most sensitive to splicing inhibition. Moreover, this sensitivity may vary depending on the mode of inhibition (Olthof et al., 2021). Minor intron-containing genes, which are targets of the minor spliceosome and make up ∼2% of all genes, are a relatively well-defined population (Olthof et al., 2019). It has been reported that the expression and splicing of minor intron-containing genes varies between tissues, and that they are least abundant in the heart and liver (Olthof et al., 2019). Notably, these two tissues are also relatively unaffected by mutations in minor spliceosome-specific components, underscoring the potential for tissue-restricted target expression to contribute to tissue specificity (Fig. 3).

Cell-type specific differences in downstream pathways could be affected by splicing inhibition

Finally, tissue-specific phenotypes might stem from cell-type specific differences in downstream signaling pathways (Fig. 5). Although loss of different spliceosome components affects the expression and function of a wide range of genes, these defects frequently converge on the activation of p53 (Allende-Vega et al., 2013; Baumgartner et al., 2018; Kleinridders et al., 2009; Lei et al., 2017; McElderry et al., 2019; White et al., 2021; Yu et al., 2019). One way by which spliceosomal defects have been shown to directly lead to p53 activation is through aberrant splicing of p53 inhibitors, such as Mdm2 and Mdm4 (Beauchamp et al., 2021; Yu et al., 2019; Alam et al., 2022). Another mode is through the formation of R-loops and induction of DNA damage (Goulielmaki et al., 2021; Sorrells et al., 2018). Cells with DNA damage also have upregulated levels of p53 in several spliceosome loss-of-function animal models, suggesting that secondary activation of p53 signaling might play a role in spliceosomopathy etiology. The fact that concomitant inhibition of p53 in spliceosome loss-of-function animal models often leads to a (partial) rescue of phenotypes such as microcephaly suggests that many developmental defects might be, in part, the result of p53 activation (Beauchamp et al., 2021; Kleinridders et al., 2009; White et al., 2021; Yu et al., 2019). Given that p53 activation itself results in cell type-specific phenotypes (Bowen et al., 2019), it is an intriguing possibility that the tissue-restricted phenotypes observed after spliceosome inhibition are the consequence of p53 activation.

In summary, splicing serves a crucial spatiotemporal regulatory role in vertebrate development. Although the significance of splicing in development is coming into focus, more studies are clearly required to deconstruct the true impact of splicing on gene expression and development. Further studies are also needed to better understand, in the context of human disease, why some cells and tissues are more affected by loss of spliceosome components than others. With advances in modern transcriptomic tools, we believe that the present is an ideal time to explore the role of splicing in development and disease.

We thank current and former members of the Kanadia lab for helpful discussions and comments on the manuscript.

Funding

The authors’ research is funded by the National Institute of Neurological Disorders and Stroke (R01NS102538). Deposited in PMC for release after 12 months.

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Competing interests

The authors declare no competing or financial interests.

Supplementary information