Neural Tube Formation and Patterning PDF

Neural Tube Formation and Patterning 13 “LIKE THE ENTOMOLOGIST IN SEARCH of brightly colored butterflies, my attention hunted, in the garden of the gray matter, cells with delicate and elegant forms, the myst...

Neural Tube Formation and Patterning 13 “LIKE THE ENTOMOLOGIST IN SEARCH of brightly colored butterflies, my attention hunted, in the garden of the gray matter, cells with delicate and elegant forms, the mysterious butterflies of the soul.” Thus reflected Santiago Ramón y Cajal, often referred to as the father of neuroscience, on his study of the brain. His 1937 quotation masterfully captures the fascination and mystery of the brain as part of a larger system that controls communication, consciousness, memory, emotion, motor control, digestion, sensory perceptions, and so much more. How the development of this central organ is coordinated with the development of the rest of the organism for integrated connectivity will remain one of the most fundamental questions in developmental biology for the next century. The first pivotal event is the transformation of an epithelial sheet into a tube. This initial structure will provide the foundation for the regionalization and diversification of brain structures along the anterior-to-posterior axis; then, through strategic mechanisms of cell growth and differentiation, the elaborate and highly connected structure of the vertebrate central nervous system can be realized. Over the next three chapters, we will study the development of the nervous system, beginning in this chapter with the formation of the neural tube and the specification of cell fates within it (FIGURE 13.1). In Chapter 14, we will delve into the mechanisms governing cell fate patterning and neurogenesis along the dorsal-ventral axis of the central nervous system. Then, in Chapter 15, we will navigate the molecular guidance mechanisms underlying the wiring of the nervous system and the development of neural crest lineages. What’s preventing this brain from closing? Courtesy of Lee Niswander and Huili Li The Punchline The vertebrate brain and spinal cord start their development as a flat plate of neuroepithelial cells that folds up and seals along most of its length to form a tube. The process of forming this neural tube is called neurulation. Folding occurs at specific locations in the plate through asymmetric changes in the shapes of cells, such that their apical sides contract, establishing hinge points of tissue bending. The folding brings the sides of the plate upward and toward each other until they fuse along the midline, as if the tube were being zipped up. The tube separates from the surface ectoderm via differential adhesion, and the central nervous system is born. The cells of the new neural tube become specialized as precursors of neurons and glial cells, and the different regions of the tube become specified along its anteroposterior and dorsoventral axes. Morphogen gradients emanating from the dorsal surface ectoderm and the ventral notochord establish positional cues for the induction of key regulatory transcription factors, which initiate cell type- specific gene regulatory networks. TGF-β and Sonic hedgehog signals play important roles in both neurulation and cell fate patterning of the neural tube. FIGURE 13.1 The major questions to be addressed in Chapters 13, 14, and 15. Questions of neurulation and cell fate specification (A,B) will be answered in this chapter. How the neural tube (NT) is expanded into the elaborate structures of the brain (C) will be covered in Chapter 14. How the peripheral nervous system (D) is largely derived from neural crest cells (NCC) migrating out of the dorsal neural tube, and how the newly born neurons extend long processes to find their synaptic partners (E) and thus connect up the nervous system will be covered in Chapter 15. The vertebrate ectoderm, the outer germ layer covering the late-stage gastrula, has three major responsibilities (FIGURE 13.2): 1. One part of the ectoderm will become the neural plate, the presumptive neural tissue induced by the prechordal plate and notochord during gastrulation. The neural plate moves into the body to form the neural tube, the precursor of the central nervous system (CNS)—the brain and spinal cord. 2. Another part of this germ layer will become the epidermis, the outer layer of the skin (which is the largest organ of the vertebrate body). The epidermis forms an elastic, waterproof, and constantly regenerating barrier between the organism and the outside world. 3. Between the compartments forming the epidermis and the central nervous system lies the presumptive neural crest. The cells of the neural crest delaminate from these epithelia at the dorsal midline and migrate away (between the neural tube and epidermis) to generate, among other things, the peripheral nervous system (all the nerves and neurons lying outside the CNS) and pigment cells (e.g., melanocytes). FIGURE 13.2 Major derivatives of the ectoderm germ layer. The ectoderm is divided into three major domains: the surface ectoderm (primarily epidermis), the neural crest (peripheral neurons, pigment, facial cartilage), and the neural tube (brain and spinal cord). The processes by which the three ectodermal regions are made physically and functionally distinct from one another is called neurulation, and an embryo undergoing these processes is called a neurula (FIGURE 13.3; Gallera 1971). As we saw in the preceding chapters, the specification of the ectoderm is accomplished during gastrulation, primarily by regulating the levels of BMP experienced by the ectodermal cells. High levels of BMP specify the cells to become epidermis. Very low levels specify the cells to become neural plate. Intermediate levels effect the formation of the neural crest cells. Neurulation directly follows gastrulation. FIGURE 13.3 Two views of primary neurulation in an amphibian embryo, showing early (left), middle (center), and late (right) neurulae in each case. (A) Looking down on the dorsal surface of the whole embryo. (B) Transverse section through the center of the embryo. (After B. I. Balinsky. 1981. Introduction to Embryology, 5th Ed. Saunders: Philadelphia.) Transforming the Neural Plate into a Tube: The Birth of the Central Nervous System The cells of the neural plate are characterized by expression of the Sox family of transcription factors (Sox1, 2, and 3). These factors (1) activate the genes that specify cells to be neural plate and (2) inhibit the formation of epidermis and neural crest by blocking the transcription and signaling of BMPs (Archer et al. 2011). In this process, we see once again an important principle of development: often the signals promoting the specification of one cell type also block the specification of an alternative cell type. The expression of Sox transcription factors establishes the neural plate cells as neural precursors that can form all the cell types of the central nervous system (Wilson and Edlund 2001). Although the neural plate lies on the surface of the embryo, the nervous system will not lie on the outside of the mature body. Somehow, the neural plate has to move inside the embryo and form a neural tube. This process is accomplished through neurulation, which occurs with some diversity across vertebrates (Harrington et al. 2009). There are two principal modes of neurulation. In primary neurulation, the cells surrounding the neural plate direct the neural plate cells to proliferate, invaginate into the body, and separate from the surface ectoderm to form an underlying hollow tube. In secondary neurulation, the neural tube arises from the aggregation of mesenchyme cells into a solid cord that subsequently forms cavities that coalesce to create a hollow tube. In many vertebrates, primary and secondary neurulation are divided spatially in the embryo such that primary neurulation forms the anterior portion of the neural tube and the posterior portion of the neural tube is the product of secondary neurulation (FIGURE 13.4). In birds, primary neurulation generates the neural tube anterior to the hindlimbs (Pasteels 1937; Catala et al. 1996). In mammals, secondary neurulation begins at the level of the sacral vertebrae of the tail (Schoenwolf 1984; Nievelstein et al. 1993). In fish and amphibians (e.g., zebrafish and Xenopus), only the tail neural tube is derived from secondary neurulation (Gont et al. 1993; Lowery and Sive 2004). More basal chordates, such as Amphioxus and Ciona, only exhibit mechanisms of primary neurulation, suggesting that primary neurulation was the ancestral condition and that secondary neurulation evolved much the way limbs did—that is, as a vertebrate novelty—and in the case of secondary neurulation, one that was associated with tail elongation (Handrigan 2003). FIGURE 13.4 Primary and secondary neurulation and the transition zone between them. The bottom image is a lateral view of the neural tube surface. The illustrations above the neural tube correspond to transverse sections through the axial level indicated as the neural tube forms in a rostral-to-caudal direction. Different cell types are represented in different colors, as indicated in the key. (After A. Dady et al. 2014. J Neurosci 34: 13208–13221.) The neural tube is finally complete when these two separately formed tubes join together (Harrington et al. 2009). The size of the transition zone between the primary and secondary neural tubes varies among species, from relatively abrupt in the mouse, to a region spanning the thoracic vertebrae in the chick, to the thoracolumbar region in humans (Dady et al. 2014). Formation of the neural tube in this transition zone has been named junctional neurulation (Dady et al. 2014) because it involves a combination of mechanisms involved in both primary and secondary neurulation (see Figure 13.4). DEV TUTORIAL 13.1 Neurulation Dr. Michael Barresi describes the cellular events and molecular mechanisms behind neural tube formation.  ? Developing Questions Why is there a need for two separate mechanisms to complete the neural tube? What were the evolutionary pressures that forced the adoption of secondary neurulation as opposed to a posterior extension of primary neurulation? As you ponder these questions, consider the embryo’s first morphogenetic mechanism, that of gastrulation. Does the timing of the end of gastrulation and its capacity for axis elongation (or lack thereof) influence your ideas? It is surprising that we still do not fully understand the evolutionary history of the nervous system. Primary neurulation Although some species differences exist, the process of primary neurulation is relatively similar in all vertebrates.1 To explore the mechanisms of neural plate folding, we will largely focus on the process of primary neurulation in amniotes. Shortly after the neural plate has formed in the chick, its edges thicken and move upward to form the neural folds, and a U-shaped neural groove appears in the center of the plate, dividing the future right and left sides of the embryo (FIGURE 13.5). The neural folds on the lateral sides of the neural plate migrate toward the midline of the embryo, eventually fusing to form the neural tube beneath the overlying ectoderm. Primary neurulation can be divided into four distinct but spatially and temporally overlapping stages: FIGURE 13.5 The neurulating chick embryo (dorsal view) at about 24 hours. The cephalic (head) region has undergone neurulation, while the caudal (tail) region is still undergoing gastrulation. (After B. M. Patten. 1971. Early Embryology of the Chick, 5th Ed. McGraw-Hill: New York; A. F. Huettner. 1943. Fundamentals of Comparative Embryology of the Vertebrates. The Macmillan Company: New York.) 1. Elongation and folding of the neural plate. Cell divisions within the neural plate are preferentially in the anterior-posterior direction (often referred to as the rostral-caudal, or beak-to-tail, direction), which fuels continued axial elongation associated with gastrulation. These events occur even if the neural plate tissue is  isolated from the rest of the embryo. To roll into a neural tube, however, the presumptive epidermis is also needed (FIGURE 13.6A,B; Jacobson and Moury 1995; Moury and Schoenwolf 1995; Sausedo et al. 1997). 2. Bending of the neural plate. The bending of the neural plate involves the formation of hinge regions where the neural plate contacts surrounding tissues. In birds and mammals, the cells at the midline of the neural plate form the medial hinge point, or MHP (Schoenwolf 1991a,b; Catala et al. 1996). MHP cells are reported to be firmly anchored to the notochord beneath them and form a hinge, which enables the creation of a furrow, or neural groove, at the dorsal midline (FIGURE 13.6C). 3. Convergence of the neural folds. Shortly thereafter, two dorsolateral hinge points (DLHPs) are induced by and anchored to the surface (epidermal) ectoderm. After the initial furrowing of the neural plate, the plate bends around the hinge regions. Each hinge acts as a pivot that directs the rotation of the cells around it (Smith and Schoenwolf 1991). Continued convergence of the surface ectoderm pushes toward the midline of the embryo, providing another motive force for bending the neural plate, causing the neural folds to converge (FIGURE 13.6D; Alvarez and Schoenwolf 1992; Lawson et al. 2001). This movement of the presumptive epidermis and the anchoring of the neural plate to the underlying mesoderm may also be important for ensuring that the neural tube invaginates, folding inward, into the embryo and not outward (Schoenwolf 1991a). 4. Closure of the neural tube. The neural tube closes as the paired neural folds are brought in contact with one another at the dorsal midline. The folds adhere to each other, and the neural and surface ectoderm cells from one side fuse with their respective counterparts from the other side. During this fusion event, cells at the apex of the neural folds delaminate and become neural crest cells (FIGURE 13.6E). FIGURE 13.6 Primary neurulation: neural tube formation in the chick embryo. (A, 1a) Cells of the neural plate can be distinguished as elongated cells in the dorsal region of the ectoderm. (B, 1b) Folding begins as the medial hinge point (MHP) cells anchor to the notochord and change their shape while the presumptive epidermal cells move toward the dorsal midline. (C, 2a) The neural folds are elevated as the presumptive epidermis continues to move toward the dorsal midline. Asymmetric constriction of actin on the apical side changes cell shapes to promote MHP bending (B, C, 2b). (C) Elevated neural folds  stained to show the extracellular matrix (green) and the actin cytoskeleton (red) concentrated in the apical portions of the neural plate cells. (D, 3a) Convergence of the neural folds occurs as the cells at the dorsolateral hinge point (DLHP) become wedge- shaped and the epidermal cells push toward the center. (D, 3b) Similar apical constriction occurs at the DLHP. (E, 4) The neural folds are brought into contact with one another. The neural crest cells disperse, leaving the neural tube separate from the epidermis. (Drawings after J. L. Smith and G. C. Schoenwolf. 1997. Trends Neurosci 20: 510–517.) REGULATION OF HINGE POINTS To fold the neural plate means to bend a sheet of epithelial cells. How can a row of attached boxlike epithelial cells be bent? While in the shape of a rectangular box (i.e., epithelial), they cannot; however, if in a region of boxes the surface area of one side of each box were reduced relative to its apposing side (creating the shape of a truncated pyramid), each of these cells should introduce a displacing angle with its neighboring cells and cause the row of boxes to bend. The MHP and two DLHPs are three regions of the neural plate where such cell shape changes occur (see Figure 13.6B–D). The epithelial cells in these locations adopt a “wedge-shaped” (or truncated pyramid) morphology along the apicobasal axis, one that is wider basally than apically (Schoenwolf and Franks 1984; Schoenwolf and Smith 1990). Similar to the bottle cells that initiate invagination during gastrulation (see Figure 11.4), localized contraction of actinomyosin complexes at the apical border reduces the size of the apical half of the cell relative to the basal compartment, a process known as apical constriction. This apical constriction pairs with the basal location of nuclei to yield the wedge-shaped hinge point cells (see Figure 13.6C,D; Smith and Schoenwolf 1987, 1988). In addition, recent findings suggest that the division rates in the dorsolateral domains of the neural plate are significantly faster than those in the ventral regions; this increases the cell density in the neural folds and adds a force that is hypothesized to promote buckling at the DLHP (McShane et al. 2015). The physical forces exerted by different regions of the neural plate have yet to be quantified, but at the cellular level, hinge points are formed by (1) apical constriction; (2) basal thickening, with retention of the nucleus within the basal portion of cells; and (3) cell packing in the neural folds. FIGURE 13.7 Morphogen regulation of hinge point formation. BMPs are expressed by the surface ectoderm (green), Noggin is expressed in the dorsal neural folds (blue), and Shh is expressed ventrally in the notochord and floor plate (orange). The regulation of hinge points revolves around BMP as an antagonist to both DLHP and MHP formation. Shh is required for the specification of floor plate, while additional signals from the notochord induce MHP morphology. Noggin directly inhibits BMP ligands, thus alleviating BMP repression of the hinge points. The DLHPs, however, form only at the correct size and dorsal-ventral position, which is based on Noggin’s distance from inhibitory Shh gradients ascending from the floor plate. Therefore, apical constriction occurs only in those cells experiencing low enough concentrations of both BMP (MHP and DLHP) and Shh (DLHP) morphogens.  What regulates these cellular changes in the correct locations of the neural plate? Here’s the short answer: Hinge point formation appears to center around the precise control of BMP signaling. BMP inhibits MHP and DLHP formation, whereas repression of BMP by Noggin enables DLHPs to form, and Shh from the notochord and floor plate prevent precocious and ectopic hinges from forming in the neural plate (FIGURE 13.7). To further understand the signaling network in control of hinge point formation and the data to support it, see Further Development 13.1, Molecular Regulation of Hinge Point Formation, online. EVENTS OF NEURAL TUBE CLOSURE Closure of the neural tube does not occur simultaneously throughout the neural ectoderm. This phenomenon is best seen in amniote vertebrates (reptiles, birds, and mammals), whose body axis is elongated prior to neurulation. In amniotes, induction occurs in an anterior-to- posterior fashion. So, in the 24-hour chick embryo, neurulation in the cephalic (head) region is well advanced, but the caudal (tail) region of the embryo is still undergoing gastrulation (see Figure 13.5). The two open ends of the neural tube are called the anterior neuropore and the posterior neuropore. ? Developing Questions What induces medial hinge point formation? Two findings—that (1) an extra notochord can induce ectopic hinge point formation and (2) Sonic hedgehog represses the DLHP—suggest that factor(s) beyond the precise control of BMP signaling may be responsible. Could it be the early expression of Noggin in the notochord (and thus still be all about repressing BMPs)? Here is an additional fact to bear in mind: in the anteriormost region of the neural plate only an MHP forms, whereas in the posteriormost region of the neural plate only DLHPs form. Only in the central regions of the neural plate are both types of hinge points present. Why are these hinge points located in different positions along the anterior-to-posterior axis, and how is this difference regulated? In chicks, neural tube closure is initiated at the level of the future midbrain and “zips up” in both directions. By contrast, in mammals, neural tube closure is initiated at several places along the anterior-posterior axis (FIGURE 13.8). In humans, there are probably five sites of neural tube closure (see Figure 13.5B; Nakatsu et al. 2000; O’Rahilly and Muller 2002; Bassuk and Kibar 2009), and the closure mechanism may differ at each site (Rifat et al. 2010). The rostral closure site (closure site 1) is located at the junction of the spinal cord and hindbrain and appears to close, as does the chick neural tube, by zipping up the neural folds. Similarly, at closure site 2, located at the midbrain/forebrain boundary, a directional zipper-like mechanism paired with dynamic cell extension appears to be at work. At closure site 3 (the rostral forebrain), the dorsolateral hinge points appear to be fully responsible for the neural tube closure. FIGURE 13.8 Neural tube closure in the mammalian embryo. (A,B) Initiation sites for neural tube closure of mouse (A) and human (B) embryos. In addition to the three initiation sites found in mice, neural tube closure in humans also initiates at the posterior end of the hindbrain and in the lumbar region. (C) Dorsal view of a 22-day (8-somite) human embryo initiating neurulation. Both anterior and posterior neuropores are open to the amniotic fluid. (D) A 10-somite human embryo showing some of the major sites of neural tube closure (arrows). (E) Dorsal view of a 23-day neurulating human embryo with only its neuropores open. (F) Midbrain exencephaly and open spina bifida are seen in the mouse curly tail mutation, a hypomorphic mutation in the grainyhead-like3 gene. (G) Vitoria de Cristo, who lived with anencephaly for two and a half years (professors, see also case study associated with this chapter). Anencephaly results when a failure to close the neural tube at sites 2 and 3 allows the forebrain to remain in contact with amniotic fluid and subsequently degenerate. (A,B after A. G. Bassuk and Z. Kibar. 2009. Semin Pediatr Neurol 16: 101–110.) FIGURE 13.9 Neural tube closure at mouse site 2 (midbrain region; see Figure 13.8A). (A) Live imaging of a 15-somite stage embryo using a transgenic CAG:Venusmyr mouse to visualize all cell membranes. Optical dorsoventral (cross) sections seen from the top image to the bottom image show DLHP formation (curving of white line on left fold) to the point of near closure (decreasing size of double arrow). (B) Optical section through a mouse embryo as the neural folds are touching but not yet closed. The single layer of non-neural surface ectoderm (large, flattened cells; stained green) has wrapped itself around the neural ectoderm (stained blue) at the edge of the closing neural folds. (C) Dotted lines show the border between neural and non- neural ectoderm. Cellular bridges from the non-neural ectoderm connect the two juxtaposed neural folds. (D) A close-up of one of these bridges (from boxed area in C) is marked by arrowheads. How do the apices of the neural folds zip up? Are there interlocking cell membranes and some mysterious force that sequentially puts them together one at a time along the anterior-to-posterior axis? One way to better understand a process as complex as neural tube closure is to simply watch it. Rather remarkable in toto live-cell imaging has been conducted on mouse embryos in culture (Pyrgaki et al. 2010; Massarwa and Niswander 2013). During DLHP bending, dynamic cell processes extend from the juxtaposed tips of the neural folds (FIGURE 13.9). This cellular behavior is being displayed by non-neural surface ectoderm cells, which ultimately extend long filopodial processes toward the opposing fold. These filopodial extensions establish temporary “cellular bridges,” whose functions are currently unknown. (See Further Development 13.2, The Biomechanics of Neural Fold Zippering Revealed by the Ancestral Chordate, online.)  SCIENTISTS SPEAK 13.1 Listen to a web conference in which Dr. Lee Niswander discusses neural tube closure. FUSION AND SEPARATION The neural tube eventually forms a closed cylinder that separates from the surface ectoderm. This separation appears to be mediated by the expression of different cell adhesion molecules. Although the cells that will become the neural tube originally express E-cadherin, they stop producing this protein as the neural tube forms and instead synthesize N-cadherin (FIGURE 13.10A). As a result, the surface ectoderm and neural tube tissues no longer adhere to each other. If the surface ectoderm is experimentally made to express N-cadherin (by injecting N-cadherin mRNA into one cell of a two-cell Xenopus embryo), the separation of the neural tube from the presumptive epidermis is dramatically impeded (FIGURE 13.10B; Detrick et al. 1990; Fujimori et al. 1990). Loss of the gene for N-cadherin in zebrafish also results in failure to form a neural tube (Lele et al. 2002). The Grainyhead transcription factors are especially important in this process (Rifat et al. 2010; Werth et al. 2010; Pyrgaki et al. 2011). Grainyhead-like2, for instance, controls a battery of cell adhesion molecules and downregulates E-cadherin synthesis in the neural folds. Mice with mutations in Grainyhead-like2 or Grainyhead-like3 genes have severe neural tube defects, which include a split face, exencephaly, and spina bifida (see Figure 13.8F and Scientists Speak 13.1; Copp et al. 2003; Pyrgaki et al. 2011). FIGURE 13.10 Expression of N- and E-cadherin adhesion proteins during neurulation in Xenopus. (A) Normal development. In the neural plate stage, N-cadherin is seen in the neural plate, whereas E-cadherin is seen on the presumptive epidermis. Eventually, the N-cadherin-bearing neural cells separate from the E-cadherin-containing epidermal cells. (Neural crest cells express neither N- nor E-cadherin, and they disperse.) (B) No separation of the neural tube occurs when one side of the frog embryo is injected with N-cadherin mRNA so that N-cadherin is expressed in the epidermal cells as well as in the presumptive neural tube. ? Developing Questions What initiates the directionality of neural tube closure? Zipping proceeds in a posterior-to-anterior direction in the tunicate Ciona, as well as in certain closure points in mammals, yet it proceeds in opposite directions to close other regions of the mammalian brain. Moreover, are the cell forces that seem to advance the zipper in the primitive chordate Ciona conserved throughout vertebrates? NEURAL TUBE CLOSURE DEFECTS In humans, neural tube closure defects occur in about 1 in every 1000 live births. Failure to close the posterior neuropore (closure site 5; see Figure 13.8B) around day 27 of development results in a condition called spina bifida, the severity of which depends on how much of the spinal cord remains exposed. Failure to close site 2 or site 3 in the rostral neural tube keeps the anterior neuropore open, resulting in a usually lethal condition called anencephaly, in which the forebrain remains in contact with the amniotic fluid and subsequently degenerates. The fetal forebrain ceases development, and the vault of the skull fails to form (see Figure 13.8G). The failure of the entire neural tube to close over the body axis is called craniorachischisis.  FURTHER DEVELOPMENT THE GENETIC AND ENVIRONMENTAL CAUSES OF NTDs Failure to close the neural tube can result from both genetic and environmental causes (Fournier-Thibault et al. 2009; Harris and Juriloff 2010; Wilde et al. 2014). Mutations (first found in mice) in genes such as Pax3, Sonic hedgehog, Grainyhead, Tfap2, and Openbrain show that these genes are essential for the formation of the mammalian neural tube; in fact, more than 300 genes appear to be involved. Environmental factors including drugs, maternal dietary factors (such as deficiencies in cholesterol, zinc and folate, also known as folic acid or vitamin B9), diabetes, obesity, and toxins can all influence human neural tube closure. How these factors lead to neural tube defects is largely unknown. For instance, a recent report has demonstrated that zinc deficiency disrupts neural tube closure by leading to the stabilization of p53 and consequently increased apoptosis (FIGURE 13.11A; Li et al. 2018). Folic acid deficiencies, however, have been one of the leading causes of neural tube defects. An emerging idea posits that a major outcome of environmental perturbations is the modification of the embryo’s epigenome, which in turn causes transcription variability leading to neural tube defects (FIGURE 13.11B; Feil et al. 2012; Shyamasundar et al. 2013; Wilde et al. 2014). This idea is most associated with the potential downstream consequences of folic acid metabolism. FIGURE 13.11 Environmental influences on neural tube defects and the role of folic acid. (A) Dorsal view of the anterior neural tube (developing brain) of control mice and of mice treated with the zinc chelator TPEN. Zinc depletion causes a dramatic increase in apoptosis, as indicated by DNA fragmentation (TUNEL labeling, green) and cleaved Caspase3 (red). Nucleic acids are stained with Hoechst dye (blue). (B) Overview of the connection that environmental  factors are proposed to have with neural tube defects (NTD). Black arrows represent the main proposal for how environmental factors may lead to neural tube defects. Gray arrows represent other possible modes leading to NTD. (C) Simplified biochemical pathway for the metabolism of folic acid leading to epigenetic regulation through DNA methylation or histone modification. DHFR, dihydrofolate reductase; MTHFR, methylenetetrahydrofolate reductase; 5- methyl-THF, 5-methyltetrahydrofolate; SAM, S-adenosylmethionine. Although the exact role of folate remains unknown, the early use of drugs that are folic acid antagonists and were given to women led to fetuses with neural tube defects. Since then, many large- scale human studies have demonstrated clear correlations of neural tube disorders with folic acid deficiency, which is the reason dietary folic acid is not only recommended for pregnant women but also systematically fortified in foods (reviewed in Wilde et al. 2014). How folic acid deficiency leads to neural tube disorders is currently an active area of research. Folic acid is an important nutrient used for regulating DNA synthesis during cell division in the brain (Anderson et al. 2012) and is also critical in regulating DNA methylation (FIGURE 13.11C). Further evidence that epigenetic mechanisms are essential for proper neural tube development are the findings that functional manipulation of histone-modifying enzymes (acetyltransferases, deacetylases, demethylases) cause neural tube defects (Artama et al. 2005; Bu et al. 2007; Shpargel et al. 2012; Welstead et al. 2012; Murko et al. 2013). Whatever the mechanisms, it has been estimated that 25%–30% of human neural tube birth defects can be prevented if pregnant women take supplemental folate. Therefore, the U.S. Public Health Service recommends that women of childbearing age take 0.4 milligram of folate daily (Milunsky et al. 1989; Centers for Disease Control 1992; Czeizel and Dudás 1992). Secondary neurulation Secondary neurulation, which takes place in the most posterior region of the embryo during tailbud elongation, produces a neural tube through a very different process than primary neurulation (see Figure 13.4). Secondary neurulation involves the production of mesenchyme cells from the prospective ectoderm and mesoderm, followed by the condensation of these cells into a medullary cord beneath the surface ectoderm (FIGURE 13.12A,B). After this mesenchymal-epithelial transition, the central portion of this cord undergoes cavitation to form several hollow spaces, or lumens (FIGURE 13.12C); the lumens then coalesce into a single central cavity (FIGURE 13.12D; Schoenwolf and Delongo 1980). FIGURE 13.12 Secondary neurulation in the caudal region of a chick embryo. (A–D) A 25-somite chick embryo. (A) Mesenchymal cells condense to form the medullary cord at the most caudal end of the chick tailbud. (B) The medullary cord at a slightly more anterior position in the tailbud. (C) The neural tube is cavitating and the notochord is forming; note the presence