Neural Tube Closure Defects Classification Essay


How the neural tube (NT) forms is a central issue in developmental biology. Vertebrate neurulation is a complex morphogenetic process that requires the coordination of many cellular and molecular events, and is regulated by more than 300 genes in mammals (Wilde et al., 2014). Primary neurulation (see Glossary, Box 1) – the process by which the NT closes from an open neural plate (NP) – is achieved sequentially in distinct steps (Fig. 1): the NP is initially induced to differentiate, it then undergoes bending to create the neural folds, which elevate towards the dorsal midline, and finally the neural fold tips fuse to complete the NT. However, primary neurulation varies between species (see Box 2). For example, there is also a sequence of neurulation events along the body axis that involves multi-site, progressive closure (‘zippering’) in mammals, a somewhat simpler sequence of closure events in birds, and almost simultaneous closure at all axial levels in amphibia (Fig. 2). Although mammals, birds and amphibians have differences in primary neurulation all undergo ‘secondary’ neurulation (Fig. 1; see Glossary, Box 1) in an apparently similar manner. In this process, a solid cord of NT progenitor cells in the developing tail bud becomes ‘canalized’ to form a neuroepithelium (NE) surrounding a lumen, without formation or closure of neural folds (Copp et al., 2015). Neurulation in teleost fish has been likened to secondary neurulation in higher vertebrates.

Box 1. Glossary

Anencephaly: Absence of brain and skull vault due to neural tissue degeneration following failed cranial neural tube closure.

Apical constriction: Contraction of the apical side of a cell.

Closure 1: The initial neural tube closure event; at the hindbrain/cervical boundary in mice.

Convergent extension (CE): The process by which a tissue narrows along its mediolateral axis (convergence) and elongates along its anteroposterior/rostrocaudal axis (extension).

Craniorachischisis: An open neural tube defect involving the midbrain, hindbrain and entire spinal cord.

Exencephaly: Failure of neural tube closure in the developing brain; the precursor of anencephaly.

Filopodia: Spike-like cellular actin-containing cellular protrusions.

Holoprosencephaly: Severe brain defect resulting from failure of the forebrain to divide into bilateral hemispheres.

Interkinetic nuclear migration (IKNM): The movement of nuclei along the apicobasal axis, depending on the cell cycle in a pseudostratified epithelium.

Junctional exchange: Cellular partner rearrangements, such as those seen during convergent extension of epithelial intercalation, in which existing cell junctions are replaced by novel junctions with different cells.

Lamellipodia (ruffles): Sheet-like cellular actin-containing cellular protrusions.

Neuromesodermal progenitors: Bi-potential progenitor cells found in the tailbud that give rise to both neuroepithelial and mesodermal derivatives.

Primary neurulation: Formation of the neural tube by bending of the neuroepithelium to generate the neural folds that elevate and fuse in the midline.

Secondary neurulation: Formation of neural tube in the lower sacral and coccygeal regions by an internal process of epithelialization, without neural fold formation or closure.

Spina bifida or spina bifida aperta: A congenital malformation of the developing spinal cord due to failure of neural tube closure (also called myelomeningocele).

Fig. 1.

An overview of primary and secondary neurulation. (A) Schematic representation of primary neurulation involving elevation of the neural folds (left panel), followed by their bending (middle panel) and fusion (right panel). DLHP, dorsolateral hinge point; MHP, median hinge point; NC, notochord; NE, neuroepithelium; NNE, non-neural ectoderm; PM, paraxial mesoderm. (B) Secondary neurulation. Tail-bud cells condense (left panel) in the midline to form the medullary cord. The medullary cord undergoes epithelialization (middle panel) around a lumen (red) while the notochordal precursor remains solid, generating the secondary neural tube and notochord (right panel).

Box 2. Variations in neurulation between vertebrates

Neurulation varies across species, both with regard to the number of closure points, their timing and the order in which these points close. In mice, the initial neural tube (NT) closure point (closure 1; see Glossary, Box 1) is at the hindbrain/cervical boundary, with a second independent initiation point (closure 2) at the forebrain/midbrain boundary. Closure (‘zippering’) proceeds both rostrally and caudally from these sites (Fig. 2A). A third initiation point (closure 3) is located at the most rostral end of the forebrain and closure proceeds backwards from this site towards closure 2 (Fig. 2A) (Copp and Greene, 2010). These multiple closure initiation sites create three neuropores (open regions of the NT): the anterior and hindbrain neuropores in the cranial region and the posterior neuropore (PNP) in the low spinal region. Neurulation in humans appears to be slightly different: human embryos between Carnegie stages 8 and 13 display two closure initiation sites, corresponding to mouse closures 1 and 3, whereas there is no apparent equivalent to closure 2 in humans (O'Rahilly and Muller, 2002). In non-mammalian vertebrates, there is progressively more divergence in the neurulation process with increasing evolutionary separation. The chick, for example, has two points of closure initiation: at the level of the future midbrain and at the hindbrain-cervical boundary, with bi-directional zippering between the sites (Fig. 2B) (Van Straaten et al., 1996). By contrast, Xenopus embryos exhibit closure almost simultaneously along the entire body axis and in teleost fish there is no formation of neural folds at all; rather, the NP cells coalesce to form a neural keel and the NT lumen opens subsequently within this structure.

Fig. 2.

Comparative schematic summary of neurulation in different vertebrates. Key features of neurulation are shown for (A) mouse, (B) chick, (C) Xenopus and (D) zebrafish. Cross-sections of the mouse, chick and Xenopus neural tube are shown with typical appearance in anterior and posterior embryonic regions; for zebrafish, cross-sections of the midbrain after closure and during neural keel formation are shown. The arrows in A and B indicate directions of closure. Arrowheads indicate hinge points. EVL, envelope layer; NE, neuroepithelium; NNE, non-neural ectoderm.

These neurulation mechanisms involve not only the NE itself but also the surrounding tissues. Indeed, the non-neural (surface) ectoderm (NNE), mesoderm and notochord have all been implicated in regulating NT closure (NTC). Importantly, failure of the dynamic morphological changes of neurulation lead to perturbations in NTC, generating neural tube defects (NTDs, see Box 3) which are among the most common human birth defects (Greene and Copp, 2014). Thus, an understanding of the mechanisms governing NTC may ultimately assist in the development of methods for predicting and preventing NTDs.

Box 3. Human neural tube defects

Neural tube defects (NTDs) are among the commonest of birth defects, with a frequency that typically ranges between 0.5 and 2 per 1000 pregnancies (Copp et al., 2015). However, in some geographical regions, e.g. Northern China, frequencies as high as 10 per 1000 births have been reported (Li et al., 2006). Failure of fusion in the cranial region results in exencephaly, which progresses to anencephaly (see Glossary, Box 1), and affected foetuses are stillborn or die postnatally. Open NT in the spinal region leads to a variety of NTDs with the most severe being myelomeningocele (also called open spina bifida or spina bifida aperta; see Glossary, Box 1). Disability in NTDs results not from abnormal differentiation of the neural tissue but rather from its progressive degeneration through contact with the amniotic fluid, making foetal surgery to cover and protect the lesion a promising therapeutic approach (Adzick et al., 2011). In addition, there is a wide spectrum of closed (skin covered) spinal dysraphisms probably caused by defective secondary neurulation, in view of their low axial level. The recent identification of neuromesodermal progenitors (Henrique et al., 2015; see Glossary, Box 1) in the tail bud might explain the poor separation of neural and mesodermal tissues in these lesions, and the frequent association with lipomas (lipomyelomeningocele) (Finn and Walker, 2007). More than 300 genes have been identified to cause NTDs in mice (Wilde et al., 2014), whereas in humans 82 genes have been considered as genetic risk factors (Pangilinan et al., 2012). Even though the risk of an affected NTD pregnancy is increased with a previous NTD occurrence, human NTDs exhibit features of multigenic inheritance with an important role for non-genetic factors (see Box 4).

In this Review, we describe recent advances in our understanding of neurulation mechanisms, focusing on the key signalling pathways, their transcriptional control, the cellular behaviours and the resulting physical forces that need to be coordinated in a regulated spatiotemporal manner. The risk factors for human NTDs and their primary prevention with nutritional supplements, including folic acid, are briefly summarized in Box 4.

Box 4. Environmental risk factors and prevention of neural tube defects

Recognized environmental influences on the development of neural tube defects (NTDs) include factors such as maternal anticonvulsant usage in early pregnancy (Werler et al., 2011) and poor nutritional status (especially low folate intake). Indeed, peri-conceptional folic acid supplementation can prevent a proportion of NTDs, as demonstrated in clinical trials (Blom et al., 2006). Although supplementation with folic acid is recommended for women planning a pregnancy, it is only countries with mandatory fortification of food with folic acid that have seen significant reductions in NTD prevalence. Some NTDs are not preventable by folic acid, and inositol has emerged as a potential preventive agent in ‘folate non-responsive’ NTD mouse models (Burren et al., 2010; Greene and Copp, 1997). A recent pilot clinical trial reported no recurrences of NTD in highly predisposed women who took inositol peri-conceptionally (Greene et al., 2016). However, larger-scale studies are needed to establish whether inositol is effective in preventing NTDs. Inositol supplementation in mouse embryo culture also rescued NTDs induced by hyperglycemia (Baker et al., 1990) raising the possibility that inositol could be beneficial in counteracting the increased risk of NTDs in maternal diabetes (Bell et al., 2012).

The initiation of NTC: the role of CE

In all of the model species in which NT formation has been studied, an initiating process involves the mediolateral (ML) convergence and anterior-posterior (AP) (or rostro-caudal) extension of axial tissues, including the NP. This process of convergent extension (CE; see Glossary, Box 1), which is responsible for shaping the NP before closure, is classically linked with the non-canonical Wnt/planar cell polarity (PCP) pathway. Accordingly, mutants in which PCP signalling is perturbed display NTDs caused by aberrant CE (where examined). As we discuss below, the study of such PCP mutants has revealed key insights into the cellular and molecular events that control CE during NTC.

Identifying a role for the PCP pathway during NTC

The non-canonical Wnt/PCP pathway is strongly conserved from Drosophila to higher vertebrates. It involves six core PCP proteins (see Box 5) that, in Drosophila, localize proximally or distally at planar polarized cell-cell junctions, forming complexes that are required for signalling function and that act to polarize cells within the plane of a tissue (reviewed by Gray et al., 2011). The significance of PCP-dependent CE in NTC was first demonstrated in post-gastrulation Xenopus embryos (Goto and Keller, 2002; Wallingford and Harland, 2002); defective CE in the midline, following the disruption of Wnt/PCP signalling, caused the NP to be too wide for the neural folds to reach the midline and fuse. Failed CE (resulting in a wide NP and short AP axis) is also observed in mice carrying mutations in core PCP genes such as Vangl2 (Greene et al., 1998). These mice fail to undergo closure 1 (see Glossary, Box 1), resulting in craniorachischisis (see Glossary, Box 1) a severe NTD in which the midbrain, hindbrain and the entire spinal region remain open (Greene et al., 1998; Murdoch et al., 2001). Similar phenotypes with a broad and short body axis have been observed in zebrafish PCP mutants (Ciruna et al., 2006; Tawk et al., 2007). Importantly, PCP mutations have been identified in humans with NTDs (reviewed by Juriloff and Harris, 2012) and evidence of abnormal CE has been observed in human foetuses with craniorachischisis (Kirillova et al., 2000). Furthermore, while homozygotes for PCP mutations display craniorachischisis, PCP heterozygotes (e.g. Vangl2Lp/+) and compound heterozygotes (e.g. Vangl2Lp/+ with mutations in Ptk7, Sec24b or syndecan 4) can develop spina bifida (see Glossary, Box 1), indicating that later events of spinal neurulation are also under the control of PCP signalling (Escobedo et al., 2013; Lu et al., 2004; Merte et al., 2010). However, even in embryos with craniorachischisis (e.g. Vangl2Lp/Lp), the forebrain NT is generally closed, suggesting that Wnt/PCP signalling may not be limiting for NTC at the most rostral levels (Greene et al., 1998; Murdoch et al., 2003).

Box 5. The planar cell polarity pathway

The core planar cell polarity (PCP) module consists of six proteins that localize in adherens junctions: three transmembrane receptors – Frizzled (Fz), Strabismus (Van Gogh or Vangl in vertebrates) and Flamingo [known as starry night (Stan) or Celsr in vertebrates] – and three cytosolic proteins – Dishevelled (Dsh; known as Dvl in vertebrates), Prickle (Pk) and Diego (Dgo; known as diversin in vertebrates). In Drosophila, these components display an asymmetric localization, but in vertebrates their distribution has not been well characterized. Downstream signalling involves small Rho-GTPases, activation of Daam1 and c-Jun N-terminal kinase (JNK), leading to a variety of cellular responses, including cytoskeletal and (in Drosophila) transcriptional regulation (Kibar et al., 2007; Roszko et al., 2009). The Fat/Dachsous module, which has been suggested to act upstream or in parallel to the PCP pathway, comprises the atypical cadherins Fat and Dachsous (Dchs) that bind each other through their large extracellular domains, and are regulated by the cytosolic kinase Four-jointed (Fjx1). Evidence from Drosophila suggests that polarized localization of Fat and Dachsous provides a cue for directional processes such as oriented cell division (Hale and Strutt, 2015; Matis and Axelrod, 2013). Other PCP-related proteins implicated in vertebrate convergent extension include Scribble and Ptk7 (protein tyrosine kinase 7).

The Fat/Dachsous/Four-jointed pathway, which functions upstream or in parallel with core PCP signalling in Drosophila (see Box 5), has also been implicated in vertebrate NTC. Fat1−/− mice display holoprosencephaly/exencephaly (see Glossary, Box 1) depending on the genetic background (Badouel et al., 2015; Ciani et al., 2003; Saburi et al., 2012), whereas Fat4−/− mice have caudal defects with a broad spinal cord (Saburi et al., 2008). In addition, members of the Fat/Dsch/Fjx pathway has been shown to interact during cranial NT closure as Fat1−/−;Fat4−/− and Fat1−/−;Fjx1−/− double mutants both display exencephaly/anencephaly (see Glossary, Box 1) whereas Fjx1−/− single mutants have no NTDs (Badouel et al., 2015; Saburi et al., 2012). It remains to be determined whether the NTDs of these mutants arise from defective CE.

Cellular mechanisms of CE

Although CE appears essential for NTC across vertebrates, the actual cellular mechanisms of CE may vary owing to differences in NP structure between animal groups (Fig. 2). The neural ectoderm in Xenopus laevis, for example, consists of two components: a superficial layer of epithelial cells overlying a deep layer of mesenchyme-like cells (Elul et al., 1997). In zebrafish, NP organization has been variously described as single-layered, bilayered (resembling amphibians) or multilayered, probably depending on the AP level (Araya et al., 2016b; Clarke, 2009), and NP cells have a columnar, epithelial-like phenotype and lack apicobasal polarity, which is established later at the neural rod stage (Girdler et al., 2013; Hong and Brewster, 2006). By contrast, the avian and mammalian NP is a single pseudostratified, columnar epithelium in which cells have obvious apicobasal polarity with nuclei positioned at different apical-basal positions within the epithelium due to interkinetic nuclear migration (IKNM; see Glossary, Box 1) (Spear and Erickson, 2012).

The cellular mechanisms of neural CE in Xenopus involve the ML intercalation of deep neural cells to form a longer and narrower NP and NT. Cell shape change and cell division do not appear to play any role. Cell movement is cell-autonomous and is accomplished by polarized protrusions that exert traction between intercalating cells (Davidson and Keller, 1999; Elul et al., 1997; Ezin et al., 2003, 2006). In parallel, the underlying mesoderm also undergoes autonomous CE, involving the formation of mediolaterally oriented bipolar protrusions (Shih and Keller, 1992). The mesoderm and midline structures dictate the behaviour of the neural cells: in their presence, NP cells exhibit lateral monopolar protrusions directed towards the NP midline (termed the ‘notoplate’ or floor plate at later stages), whereas they display mediolaterally oriented bipolar protrusions when mesoderm or midline structures are absent (Elul et al., 1997; Ezin et al., 2003, 2006). CE in both neural and mesodermal cells is regulated by the Wnt/PCP pathway, and the perturbation of PCP/CE in the NP results in NTDs (Wallingford and Harland, 2001).

In zebrafish, the perturbation of Wnt/PCP signalling (e.g. in Vangl2 mutants) results in a rather broad and thick NE (Ciruna et al., 2006; Tawk et al., 2007) due to faulty cell intercalation (Ciruna et al., 2006). Wnt/PCP signalling seems to be important for the repolarization and integration of daughter cells in the neural keel after the transient loss of polarity during division (Ciruna et al., 2006). Moreover, mirror-image NTs have been observed when Wnt/PCP signalling is abrogated, due to divisions happening at the right time but in a late-converged NE (Tawk et al., 2007). Convergence movements of the mesoderm also play a key role during CE of the zebrafish NE (Araya et al., 2014), due to the coupling role of extracellular matrix (ECM) between the two tissues (Araya et al., 2016a), although Wnt/PCP signalling seems to be important predominantly in the NE (Ciruna et al., 2006).

In mice, axial extension at E7.5-8.5, prior to closure 1, was demonstrated by vital cell labelling and shown to depend on functional Wnt/PCP signalling (Ybot-Gonzalez et al., 2007b). Vangl2Lp/Lp embryos exhibit severely diminished midline extension, resulting in widely spaced neural folds that fail to initiate NTC (Greene et al., 1998). Chimaera analysis shows that the requirement for PCP during CE is cell-autonomous, as Vangl2Lp/Lp cells exhibit reduced midline intercalation even when intermixed with wild-type cells in the same embryo (Ybot-Gonzalez et al., 2007b). Live imaging reveals that neuroepithelial cells organize into tetrads and multicellular rosettes as the AP-oriented junctions contract (Williams et al., 2014). These rosettes then resolve and new ML junctions are formed, leading to neighbour exchange and tissue elongation (apical junctional intercalation). In parallel, cells elongate basally along the ML axis, exhibiting protrusive activity and driving cells to undergo ML intercalation. These intercalation events are adversely affected in PCP mutants and result in defective CE (Williams et al., 2014).

Molecular mechanisms controlling CE

Recent studies are beginning to provide insights into how PCP signalling functions at the molecular level during the initial stages of NTC. However, Wnt/PCP signalling interfaces with cytoskeletal remodelling and thus could also participate in the regulation of later cytoskeleton-driven events, for example NP bending.

The core PCP proteins display a highly polarized cellular distribution in Drosophila epithelia and, recently, similar cellular polarization has been described in vertebrate CE, during inner ear development in mammals (May-Simera and Kelley, 2012). In Xenopus, NP cells also display anteriorly polarized Vangl2 localization. This polarization is dependent on the formation of a complex between Vangl2 and Prickle, and Wnt-dependent phosphorylation of Vangl2, which is important for normal NTC (Ossipova et al., 2015). Moreover, Vangl2 polarization depends on Rho kinase-induced phosphorylation of myosin light chain (pMLC), and it has further been shown that Vangl2 depletion abrogates MLC phosphorylation, suggesting a feedback loop between Wnt/PCP signalling and actomyosin dynamics (Ossipova et al., 2014, 2015).

Similarly, in chick neuroepithelial cells, the Wnt/Fz-PCP component Celsr1 displays polarized localization specifically at AP junctions. Celsr1 recruits Dvl, which activates Daam1, which in turn binds and activates PDZ-RhoGEF. This likely leads to RhoA activation and myosin contraction (Fig. 3), resulting in midline convergence as well as NP bending in the ML direction (Nishimura et al., 2012). In support of this sequence was the observation that pMLC is detected only in a subpopulation of adherens junctions in the bending NP of chick embryos (Nishimura and Takeichi, 2008), whereas, in mouse, basal (but not apical) myosin IIB is enriched preferentially along the AP axis (Williams et al., 2014). This polarity is disturbed in mouse PCP mutants, consistent with the lack of ML intercalation (Williams et al., 2014).

Fig. 3.

Schematic representation of key neural tube closure regulatory mechanisms. A number of mechanisms involved in neural tube closure (NTC) are depicted. (1) Transcriptional regulation: Grhl2 (grainyhead-like 2) regulates the expression of E-cadherin and Cldn4 (claudin 4) in non-neural ectoderm (NNE) cells during mouse cranial neurulation. (2) Protrusions: NNE cells display Rac1-dependent protrusions that make the first contact during neural fold (NF) fusion in the mouse spinal region. (3) Proteases: a pathway involving membrane-bound serine proteases (e.g. protease-activated receptor 2, Par2) is active in NNE cells. (4) Interkinetic nuclear migration (IKNM): nuclei migrate apically to divide, with daughter nuclei returning to a basal position for S phase. As neuroepithelial cell cycles are not synchronized, the neural plate (NP) is a pseudostratified epithelium. (5) Dorsolateral hinge point (DLHP) regulation: the formation of DLHPs is regulated by antagonistic interactions between bone morphogenetic protein 2 (BMP2), sonic hedgehog (Shh) and Noggin. (6) BMP and transforming growth factor (TGF) signalling: active BMP (detected by pSMAD1/5/8) and TGFβ (detected by pSMAD2/3) signalling are found along the neural ectoderm (NE) in a cell-cycle dependent manner. Antagonism between the pathways is important for the formation of the median hinge point (MHP) in chick midbrain, by affecting the localization of apical (e.g. PAR3) or basolateral (e.g. lethal giant larva; LGL) junctional proteins. (7) Planar polarized actomyosin contraction: planar cell polarity (PCP)-controlled apical constriction (actin fibres in red) causes bending along the mediolateral axis in the cranial neural tube of the chick. Basal nuclear localization causes wedge-shaped cells in the midline NP of both chick and mouse embryos. (8) Actomyosin turnover and extracellular matrix (ECM): the assembly and disassembly of apical actin filaments is under ROCK/RhoA regulation. ECM proteins (e.g. fibronectin, perlecan, glypican 4) and their receptors (e.g. integrins) affect NTC.

In Xenopus, diversin, another PCP component that is required for NTC, was found to be polarized along the ML axis of the NP (Ossipova et al., 2014). Rab11, a protein that is involved in endosome recycling and is also important for NTC, was found to be polarized and regulated by diversin, with pMLC acting as a downstream effector (Ossipova et al., 2014).

The available evidence suggests, therefore, that PCP-dependent polarized protein localization in both the AP and ML axes is important for CE, NP bending and ultimately NTC. Hence, vertebrate PCP signalling appears to function similarly to Drosophila planar polarization. The downstream regulator of this polarized expression across species is the phosphorylation of MLC, which is important for both CE and bending of the NP in Xenopus, chick and mouse embryos (Nishimura et al., 2012; Rolo et al., 2009; Williams et al., 2014). It remains to be determined to what extent MLC activation is required for vertebrate neurulation because of its role in CE, as opposed to its involvement in the later events of NP bending and neural fold elevation.

Bending of the NP and elevation of the neural folds

Once CE is under way, shaping the future CNS along its AP and ML axes, the margins of the NP begin to elevate, forming neural folds that eventually meet in the dorsal midline and fuse to create the NT (Fig. 1A). Amphibia undergo neural fold bending, elevation and fusion at all axial levels over a relatively short time-frame, whereas birds and mammals display a much more protracted progression of closure along the body axis (Fig. 2). Moreover, the morphology of neural fold elevation differs between cranial and spinal regions (Fig. 2). In the mouse midbrain, the neural folds are initially biconvex with their tips orientated away from the midline. Then dorsolateral bending occurs, generating biconcave neural folds and orienting the tips towards the midline for fusion (Morriss-Kay, 1981). This biphasic sequence has been related to an initial expansion of the underlying cranial mesoderm, which causes the neural folds to adopt the biconvex morphology (Zohn and Sarkar, 2012), and to subsequent actomyosin-dependent dorsolateral bending to generate the biconcave morphology (Morriss-Kay and Tuckett, 1985). In addition, the emigration of cranial neural crest, which begins before closure (unlike in the spine where it follows closure), may enable dorsolateral bending (Copp, 2005). The mammalian spinal region, by contrast, does not exhibit a biconvex elevation phase: the neural folds remain straight except for focal bending sites at the midline (the MHP; median hinge point) and dorsolaterally (DLHPs; paired dorsolateral hinge points). Moreover, as the wave of closure progresses along the spinal region, bending shifts from being predominantly MHP mediated to mainly DLHP mediated (Shum and Copp, 1996).

Cellular mechanisms involved in NP bending

Epithelial bending during development is often represented as resulting from a generalized reduction in apical surface area (apical constriction; see Glossary, Box 1). Indeed, in Xenopus embryos, apical constriction of the superficial NP layer leads to invagination at the midline and neural fold bending along the body axis. The anterior NP region additionally exhibits paired dorsolateral bending points, in which apical constriction is particularly marked (Fig. 2C) (Haigo et al., 2003; Lee et al., 2007). The disruption of apical constriction, e.g. by depletion of Shroom3 (Haigo et al., 2003) or of GEF-H1, a RhoA-specific GEF (Itoh et al., 2014), leads to NTC failure. Recently, it was shown that apical constriction in Xenopus is the result of Ca2+-dependent asynchronous and cell-autonomous actin contractions followed by calpain 2-dependent stabilization steps (Christodoulou and Skourides, 2015). In zebrafish, although neural folds do not form per se, structures resembling DLHPs have been described as playing a role in lumen formation within the NT (Fig. 2D) (Hong and Brewster, 2006; Nyholm et al., 2009).

In birds and mammals, the NP is pseudostratified, raising the issue of what constitutes ‘apical constriction’ in such an epithelium. Neuroepithelial cell shape is determined by nuclear position, which varies from apical (during mitosis) to basal (during S-phase of the cell cycle) due to the process of IKNM (Fig. 3). In most parts of the NP, cells are randomly distributed throughout the IKNM cycle, whereas the midline contains a high proportion of S-phase cells with a wedge shape, owing to their basally located nuclei. This generates local bending at the MHP directly overlying the notochord (McShane et al., 2015; Schoenwolf and Smith, 1990). The NP of the lower spine also bends at paired DHLPs, but these show no consistent association with nuclear position or cell cycle phase (McShane et al., 2015; Schoenwolf and Franks, 1984), suggesting an alternative cellular mechanism of bending at DLHPs.

Actomyosin dynamics also appear to play a role in NP bending in amphibia, birds and mammals (Fig. 3), where actin filaments, along with pMLC and Rho GTPases, are found in the apical junctions of neuroepithelial cells (Escuin et al., 2015; Kinoshita et al., 2008; Rolo et al., 2009). Thus, the actomyosin machinery is located at the right place to act as cellular ‘purse strings’, generating the force to pull the neural folds together (Sawyer et al., 2010). Indeed, mice with mutations affecting actin-associated proteins exhibit cranial NTDs, although, with the exception of Shroom3 and MARCKS-related protein mutants, these mutants notably do not develop spinal NTDs (Copp and Greene, 2010). Moreover, inhibition of actin polymerization using cytochalasin D or latrunculin B in cultured mouse embryos does not prevent the formation of MHP and DLHPs, and NTC proceeds in the spinal region (Escuin et al., 2015; Ybot-Gonzalez and Copp, 1999). Recent work has shown that spinal closure requires ROCK-dependent disassembly of actin filaments (Fig. 3

1. Wallingford JB, Niswander LA, Shaw GM, Finnell RH. The continuing challenge of understanding, preventing, and treating neural tube defects. Science. 2013;339:1222002.[PMC free article][PubMed]

2. Copp AJ, Stanier P, Greene ND. Neural tube defects: recent advances, unsolved questions, and controversies. Lancet Neurol. 2013;12:799–810.[PMC free article][PubMed]

3. Hamamy H. Epidemiological profile of neural tube defects in Arab countries. Middle East Journal of Medical Genetics. 2014;3:1–10.

4. Bassuk AG, Kibar Z. Genetic basis of neural tube defects. Semin Pediatr Neurol. 2009;16:101–110.[PubMed]

5. Copp AJ, Greene ND. Genetics and development of neural tube defects. Pathol J. 2010;220:217–230.[PMC free article][PubMed]

6. Aguilera S, Soothill P, Denbow M, Pople I. Prognosis of spina bifida in the era of prenatal diagnosis and termination of pregnancy. Fetal Diagn Ther. 2009;26:68–74.[PubMed]

7. Roebroeck ME, Jahnsen R, Carona C, Kent RM, Chamberlain MA. Adult outcomes and lifespan issues for people with childhood-onset physical disability. Dev Med Child Neurol. 2009;51:670–678.[PubMed]

8. Golden JA, Chernoff GF. Multiple sites of anterior neural tube closure in humans: evidence from anterior neural tube defects (anencephaly) Pediatrics. 1995;95:506–510.[PubMed]

9. Olney RS, Mulinare J. Trends in neural tube defect prevalence, folic acid fortification, and vitamin supplement use. Semin Perinatol. 2002;26:277–285.[PubMed]

10. Siffel C, Wong LY, Olney RS, Correa A. Survival of infants diagnosed with encephalocele in Atlanta 1979-98. Paediatr Perinat Epidemiol. 2003;17:40–48.[PubMed]

11. Tubbs RS, Wellons JC, 3rd, Oakes WJ. Occipital encephalocele, lipomeningomyelocele, and Chiari I malformation: case report and review of the literature. Childs Nerv Syst. 2003;19:50–53.[PubMed]

12. Brown MS, Sheridan-Pereira M. Outlook for the child with a cephalocele. Pediatrics. 1992;90:914–919.[PubMed]

13. Jones KL, editor. Philadelphia (PA): Elsevier Saunders; 2006. Smith's recognizable patterns of human malformation.

14. Kheir AEM, Imam A, Omer IM, Hassan IMA, Elamin SA, Awadalla EA, et al. Meckel-Gruber Syndrome: A rare and lethal anomaly. Sudan J Paediatr. 2012;12:93–96.

15. Mohamed S, Ibrahim F, Kamil K, Satti SA. Meckel-Gruber syndrome: Antenatal diagnosis and ethical perspectives. Sudan J Paediatr. 2012;12:70–72.

16. Seidahmed MZ, Abdelbasit OB, Shaheed MM, Alhussein KA, Miqdad AM, Samadi AS, et al. Genetic, chromosomal, and syndromic causes of neural tube defects. Saudi Med J. 2014;35(Suppl 1):S49–S56.[PMC free article][PubMed]

17. Balci S, Tekşen F, Dökmeci F, Cengiz B, Cömert RB, Can B, et al. Prenatal diagnosis of Meckel-Gruber syndrome and Dandy-Walker malformation in four consecutive affected siblings, with the fourth one being diagnosed prenatally at 22 weeks of gestation. Turk J Pediatr. 2004;46:283–288.[PubMed]

18. Chen CP. Meckel syndrome: genetics, perinatal findings, and differential diagnosis. Taiwan J Obstet Gynecol. 2007;46:9–14.[PubMed]

19. Salonen R, Norio R. The Meckel syndrome in Finland: epidemiologic and genetic aspects. Am J Med Genet. 1984;18:691–698.[PubMed]

20. Teebi AS, Teebi SA. Genetic diversity among the Arabs. Community Genet. 2005;8:21–26.[PubMed]

21. Parelkar SV, Kapadnis SP, Sanghvi BV, Joshi PB, Mundada D, Oak SN. Meckel-Gruber syndrome: A rare and lethal anomaly with review of literature. J Pediatr Neurosci. 2013;8:154–157.[PMC free article][PubMed]

22. Chen CP. Syndromes, disorders and maternal risk factors associated with neural tube defects (III) Taiwan J Obstet Gynecol. 2008;47:131–140.[PubMed]

23. Valente EM, Logan CV, Mougou-Zerelli S, Lee JH, Silhavy JL, Brancati F, et al. Mutations in TMEM216 perturb ciliogenesis and cause Joubert, Meckel and related syndromes. Nat Genet. 2010;42:619–625.[PMC free article][PubMed]

24. Shaheen R, Faqeih E, Seidahmed MZ, Sunker A, Alali FE, AlQahtani K, et al. A TCTN2 mutation defines a novel Meckel Gruber syndrome locus. Hum Mutat. 2011;32:573–578.[PubMed]

25. Shaheen R, Faqeih E, Alshammari MJ, Swaid A, Al-Gazali L, Mardawi E, et al. Genomic analysis of Meckel-Gruber syndrome in Arabs reveals marked genetic heterogeneity and novel candidate genes. Hum Genet. 2013;21:762–768.[PMC free article][PubMed]

26. Joubert M, Eisenring JJ, Robb JP, Andermann F. Familial agenesis of the cerebellar vermis. A syndrome of episodic hyperpnea, abnormal eye movements, ataxia, and retardation. Neurology. 1969;19:813–825.[PubMed]

27. Badano JL, Mitsuma N, Beales PL, Katsanis N. The ciliopathies: an emerging class of human genetic disorders. Annu Rev Genomics Hum Genet. 2006;7:125–148.[PubMed]

28. Khan AO, Oystreck DT, Seidahmed MZ, AlDrees A, Elmalik SA, Alorainy IA, et al. Ophthalmic features of Joubert syndrome. Ophthalmology. 2008;115:2286–2289.[PubMed]

29. Alorainy IA, Sabir S, Seidahmed MZ, Farooqu HA, Salih MA. Brain stem and cerebellar findings in Joubert syndrome. J Comput Assist Tomogr. 2006;30:116–121.[PubMed]

30. Thomas S, Wright KJ, Le Corre S, Micalizzi A, Romani M, Abhyankar A, et al. A homozygous PDE6D mutation in Joubert syndrome impairs targeting of farnesylated INPP5E protein to the primary cilium. Hum Mutat. 2014;35:137–146.[PMC free article][PubMed]

31. Alazami AM, Alshammari MJ, Salih MA, Alzahrani F, Hijazi H, Seidahmed MZ, et al. Molecular characterization of Joubert syndrome in Saudi Arabia. Hum Mutat. 2012;33:1423–1428.[PubMed]

32. Aicardi J. 3rd ed. London (UK): Mac Keith Press; 2009. Diseases of the Nervous System in Childhood.

33. Tortori-Donati P, Rossi A. Congenital malformations of the spine and spinal cord. Rivista di Neuroradiologia. 2004;17:249–267.

34. Mirsky DM, Schwartz ES, Zarnow DM. Diagnostic features of myelomeningocele: The role of ultrafast fetal MRI. Fetal Diagn Ther. 2014 Jul 22; Epub ahead of print. [PubMed]

35. Matson DD. 2nd ed. Springfield (IL): Charles C. Thomas; 1969. Neurosurgery of Infancy and Childhood.

36. Thompson DN. Postnatal management and outcome for neural tube defects including spina bifida and encephalocoeles. Prenat Diagn. 2009;29:412–419.[PubMed]

37. Elgamal EA. Natural history of hydrocephalus in children with spinal open neural tube defect. Surg Neurol Int. 2012;3:112.[PMC free article][PubMed]

38. Cama A, Tortori-Donati P, Piatelli GL, Fondelli MP, Andreussi L. Chiari complex in children--neuroradiological diagnosis, neurosurgical treatment and proposal of a new classification (312 cases) Eur J Pediatr Surg. 1995;5(Suppl 1):35–38.[PubMed]

39. Tortori-Donati P, Rossi A, Cama A. Spinal dysraphism: a review of neuroradiological features with embryological correlations and proposal for a new classification. Neuroradiology. 2000;42:471–491.[PubMed]

40. McLone DG, Knepper PA. The cause of Chiari II malformation: a unified theory. Pediatr Neurosci. 1989;15:1–12.[PubMed]

41. Naidich TP, McLone DG, Fulling KH. The Chiari II malformation: Part IV The hindbrain deformity. Neuroradiology. 1983;25:179–197.[PubMed]

42. Ellenbogen RG. Neural tube defects in the neonatal period. [Accessed 2014 March 2]. Available from: .

43. Naim-Ur-Rahman Salih MA, Jamjoom AH, Jamjoom ZA. Congenital intramedullary lipoma of the dorsocervical spinal cord with intracranial extension: case report. Neurosurgery. 1994;34:1081–1083.[PubMed]

44. Khoshhal KI, Murshid WR, Elgamal EA, Salih MA. Tethered cord syndrome: A study of 35 patients. Journal of Taibah University Medical Sciences. 2012;7:23–28.

45. Elgamal EA, Hassan HH, Elwatidy SM, Altwijri I, Alhabib AF, Jamjoom ZB, et al. Split cord malformation associated with spinal open neural tube defect. Saudi Med J. 2014;35(Suppl 1):pS44–S48.[PMC free article][PubMed]

46. Finer NN, Bowen P, Dunbar LG. Caudal regression anomalad (sacral agenesis) in siblings. Clin Genet. 1978;13:353–358.[PubMed]

47. Merello E, De Marco P, Ravegnani M, Riccipetitoni G, Cama A, Capra V. Novel MNX1 mutations and clinical analysis of familial and sporadic Currarino cases. Eur J Med Genet. 2013;56:648–654.[PubMed]

48. Lynch SA, Wang Y, Strachman T, Bum J, Lindsay S. Autosomal dominant sacral agenesis: Currarino syndrome. J Med Genet. 2000;37:561–566.[PMC free article][PubMed]

49. Tortori-Donati P, Fondelli MP, Rossi A, Raybaud CA, Cama A, Capra V. Segmental spinal dysgenesis: neuroradiologic findings with clinical and embryologic correlation. AJNR Am J Neuroradiol. 1999;20:445–456.[PubMed]

50. Lynch SA. Non-multifactorial neural tube defects. Am J Med Genet C Semin Med Genet. 2005;135:69–76.[PubMed]

51. Manning SM, Jennings R, Madsen JR. Pathophysiology, prevention, and potential treatment of neural tube defects. Ment Retard Dev Disabil Res Rev. 2000;6:6–14.[PubMed]

52. Harris MJ, Juriloff DM. Mouse mutants with neural tube closure defects and their role in understanding human neural tube defects. Birth Defects Res A Clin Mol Teratol. 2007;79:187–210.[PubMed]

53. Detrait ER, George TM, Etchevers HC, Gilbert JR, Vekemans M, Speer MC. Human neural tube defects: developmental biology, epidemiology, and genetics. Neurotoxicol Teratol. 2005;27:515–524.[PMC free article][PubMed]

54. Kennedy D, Chitayat D, Winsor EJ, Silver M, Toi A. Prenatally diagnosed neural tube defects: ultrasound, chromosome, and autopsy or postnatal findings in 212 cases. Am J Med Genet. 1998;77:317–321.[PubMed]

55. Chen CP. Chromosomal abnormalities associated with neural tube defects (I): full aneuploidy. Taiwan J Obstet Gynecol. 2007;46:325–335.[PubMed]

56. Chen CP. Chromosomal abnormalities associated with neural tube defects (II): partial aneuploidy. Taiwan J Obstet Gynecol. 2007;46:336–351.[PubMed]

57. Chen CP. Syndromes, disorders and maternal risk factors associated with neural tube defects (I) Taiwan J Obstet Gynecol. 2008;47:1–9.[PubMed]

58. Chen CP. Syndromes, disorders and maternal risk factors associated with neural tube defects (II) Taiwan J Obstet Gynecol. 2008;47:10–17.[PubMed]

59. Rampersaud E, Bassuk AG, Enterline DS, George TM, Siegel DG, Melvin EC, et al. Whole genomewide linkage screen for neural tube defects reveals regions of interest on chromosomes 7 and 10. J Med Genet. 2005;42:940–946.[PMC free article][PubMed]

60. Stamm DS, Rampersaud E, Slifer SH, Mehltretter L, Siegel DG, Xie J, et al. High-density single nucleotide polymorphism screen in a large multiplex neural tube defect family refines linkage to loci at 7p21.1-pter and 2q33.1-q35. Birth Defects Res A Clin Mol Teratol. 2006;76:499–505.[PMC free article][PubMed]

61. Stamm DS, Siegel DG, Mehltretter L, Connelly JJ, Trott A, Ellis N, et al. Refinement of 2q and 7p loci in a large multiplex NTD family. Birth Defects Res A Clin Mol Teratol. 2008;82:441–452.[PubMed]

62. Mills JL, Scott JM, Kirke PN, McPartlin JM, Conley MR, Weir DG, et al. Homocysteine and neural tube defects. J Nutr. 1996;126:756–760.[PubMed]

63. Beaudin AE, Stover PJ. Insights into metabolic mechanisms underlying folate-responsive neural tube defects: a minireview. Birth Defects Res A Clin Mol Teratol. 2009;85:274–284.[PMC free article][PubMed]

64. Gos M, Sliwerska E, Szpecht-Potocka A. Mutation incidence in folate metabolism genes and regulatory genes in Polish families with neural tube defects. J Appl Genet. 2004;45:363–368.[PubMed]

65. Frosst P, Blom HJ, Milos R, Goyette P, Sheppard CA, Matthews RG, et al. A candidate genetic risk factor for vascular disease: a common mutation in methylenetetrahydrofolate reductase. Nat Genet. 1995;10:111–113.[PubMed]

66. Ou CY, Stevenson RE, Brown VK, Schwartz CE, Allen WP, Khoury MJ, et al. 5,10 Methylenetetrahydrofolate reductase genetic polymorphism as a risk factor for neural tube defects. Am J Med Genet. 1996;63:610–614.[PubMed]

67. Relton CL, Wilding CS, Pearce MS, Laffling AJ, Jonas PA, Lynch SA, et al. Gene-gene interaction in folate-related genes and risk of neural tube defects in a UK population. J Med Genet. 2004;41:256–260.[PMC free article][PubMed]

68. Botto LD, Yang Q. 5,10-Methylenetetrahydrofolate reductase gene variants and congenital anomalies: a HuGE review. Am J Epidemiol. 2000;151:862–877.[PubMed]

69. van der Put NM, Eskes TK, Blom HJ. Is the common 677C-->T mutation in the methylenetetrahydrofolate reductase gene a risk factor for neural tube defects? A meta-analysis. QJM. 1997;90:111–115.[PubMed]

70. Wilcken B, Bamforth F, Li Z, Zhu H, Ritvanen A, Renlund M, et al. Geographical and ethnic variation of the 677C>T allele of 5,10 methylenetetrahydrofolate reductase (MTHFR): findings from over 7000 newborns from 16 areas world wide. J Med Genet. 2003;40:619–625.[PMC free article][PubMed]

71. van der Linden IJ, Afman LA, Heil SG, Blom HJ. Genetic variation in genes of folate metabolism and neural-tube defect risk. Proc Nutr Soc. 2006;65:204–215.[PubMed]

72. Blom HJ, Shaw GM, den Heijer M, Finnell RH. Neural tube defects and folate: case far from closed. Nat Rev Neurosci. 2006;7:724–731.[PMC free article][PubMed]

73. Copp AJ, Green ND. Genetics and development of neural tube defects. J Pathol. 2010;220:217–230.[PMC free article][PubMed]

74. Zhang T, Lou J, Zhong R, Wu J, Zou L, Sun Y, et al. Genetic variants in the folate pathway and the risk of neural tube defects: a meta-analysis of the published literature. PLoS One. 2013;8:e59570.[PMC free article][PubMed]


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