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3 Where Magic Happens: Development of the Embryo
Neurulation follows two stages: primary and secondary. Primary neurulation71
begins after gastrulation when the primitive ectoderm is induced by the axial mesoderm to form a neural plate. The neural plate undergoes further elevation, folding,
and fusion to form the neural tube. Neural crest cells migrate from the dorsal aspect
of the neural tube. Primary neurulation forms all functional levels of the brain and
spinal cord to the second sacral level in humans.
The caudal elements of the spinal cord, conus medullaris and filum terminale,
are formed by secondary neurulation,72–78 which begins at a transitional zone where
the dorsally located primary neural tube overlaps the more ventral mesenchymal cells
of the tail bud in the future lumbosacral area. In this overlap zone, randomly arranged
mesenchymal cells condense to form the medullary cord. Radially oriented peripheral
cells surround a cellular central core in the medullary cord. Cavitation occurs centrally,
forming multiple lumina that coalesce to form a secondary neural tube.
The source of secondary neural tube cells is under scrutiny. Recent evidence in
chick embryos suggests that cells may migrate from more rostral neural plates to
attain their proper positions in the secondary neural tubes.79,80 Normal caudal spinal
cord patterning in humans has been described81 and abnormal patterning has been
demonstrated in dysraphic states.82,83 Aberrant positional identity of caudal spinal
cord cells may be a consequence of disrupted positional signals, faulty differentiation, or improper migration. Governing factors in the caudal neural tube pattern such
as the brachyury and Pax-3 patterning genes have not been identified as major factors
in spinal dysraphism.54
10.4 MAGIC PILLS
Exciting and provocative evidence demonstrates that some manifestations of NTDs
are preventable or reversible at any one of numerous steps along the pathway from
preconception to childhood, and possibly even into adulthood (Figure 10.1). Several
different therapeutic interventions (or “magic pills”) may be developed to treat the
remaining types of NTDs. These pills may target genetic loci, proteins, or any of
several metabolites involved in NTD development.
We now understand a great deal about the development of the neural tube, and
are quickly approaching a more complete genetic characterization of the process.
Ideally, NTDs could be detected early enough in development to target the defects
before any permanent manifestations occurred. The epidemiological studies
described definitively implicate maternal risk factors as well as inheritable and/or
acquired genetic influences that may be targeted. The combination of genetic, epigenetic, and environmental factors offers numerous targets for interventions.
Preconception would be the optimal time for prevention. Mothers with modifiable risk factors should be identified and counseled. Perhaps one of the most remarkable advances in NTD treatment has been the introduction of periconceptional folic
acid supplementation for the prevention of myelodysplasias. Whether taken in pill
form or supplemented in dietary flour, this simple and inexpensive measure has cut
the incidence and devastating sequelae of myelomeningocele by more than half.
Despite this extraordinary achievement, it is still a challenge to prevent this unfortunate disorder of aberrant neural tube closure.
© 2005 by CRC Press LLC
FIGURE 10.1 The magic phases of spinal dysraphism.
Other maternal risk factors that may prove important include good control of
diabetes, reduction of obesity and infections, vitamin supplementation (folate, inositol, and vitamin B12), and avoidance of over-heated environments like saunas.
Additionally, mothers taking valproate and carbamazapine antiepileptic medications
should discontinue use or take other medications if possible to eliminate the
It may be possible in some cases to identify mothers with inheritable genetic
predispositions and counsel them during the preconception period in preparation for
possible treatment during pregnancy. Several possible medications could be developed to provide genetic targeting during early fetal development. Tools for targeting
candidate genes at the DNA, RNA, or protein level are all plausible possibilities.
These tools could target defects in genes involved in proper neural tube patterning,
folate-dependent and -independent mechanisms, or healing mechanisms. The next
decade certainly will see attempts at in vitro correction of genetic defects during the
blastocyst stage or manipulation of these genes in utero via delivery systems like
Several studies with animal models have elucidated some of the genes involved
in the induction of proper neural tube development, for example, Wnt-1, Gnot1 (a
notochord family homeobox gene), HOX-1, and activin.50,56,60 Activin and retinoic
acid regulate Gnot1 expression prior to gastrulation. The neural tube-inducing properties of sonic and bone morphogenic protein genes are also under intense investigation. The Sp mouse model has defects in neural tube closure due to mutations in
the Pax-3 paired box gene.44,45 When genes are deleted or mutated, the fetal cells
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may be transfected in utero with viral vectors expressing the normal gene. Alternatively, embryonic stem cell lines with normal genes may be introduced into target
embryos by blastocyst injection, producing chimeras expressing enough of the normal gene to ameliorate the defective phenotype. Interestingly, folate, the earliest
magic pill, has been shown to prevent NTD in the Sp and other mouse models with
mutations in Cart1 and crooked tail genes.67,84,85
Hyperhomocysteinemia is another risk factor linked to increased risk of NTD
that may be amenable to a genetic tool. The condition appears to be due to homozygosity of a thermolabile MTHFR deficiency.20 Genetic therapy could provide a
solution. Currently available viral vectors could be designed to transfect fetal cells
with the normal MTHFR gene. Hyperhomocysteinemia may also be due to reduced
folate-dependent homocysteine remethylation, which provides another interesting
mechanism for treating NTD.
Cytosine methylation on CpG dinucleotides of genomic DNA is one of many
forms of DNA modifications that help maintain stability of numerous regions of
genomic DNA.86 These heritable CpG methylation sites may be altered in early
embryogenesis, but appear to remain stable with high fidelity afterward.87 This form
of DNA methylation depends on the synthesis of S-adenosylmethionine, which
requires methyl donors and cofactors like folate, vitamin B12, choline chloride, and
Maternal nutrition may affect fetal phenotype via DNA methylation. The areas
of methylation that change during embryogenesis are at transposable element insertion sites in the genome that underlie epigenetic-induced phenotypic variability.89
Transient exposure to methyl donors in utero has been demonstrated to shift an
epigenotype via CpG methylation of genomic DNA in mice.19 This experimentally
altered phenotype persisted into adulthood. It is hypothesized that such a mechanism
may underlie the corrected NTD phenotype in folate supplementation. Other methyl
donors may also serve as magic pills.
Another compound that prevented folate resistance NTD in the curly tail mouse
and recently in humans is inositol.90,91 The mechanism may occur via upregulation
of the retinoic acid receptor beta.91,92 Inositol is also important in glucose metabolism
and may play a role in hyperglycemic or obesity-related causes of NTD. All these
therapeutic measures are meant to prevent or correct defects early enough in development to prevent NTDs. However, efforts to correct defects are still needed. Most
forms of what we can designate as “magic repairs” are applied during intrauterine
development or after birth.
In a typical scenario, a child born with a NTD undergoes repair of the defect in the
first few days after birth (as with myelomeningocele) or when neurological deterioration or substantial neurological risk is determined (as with closed dysraphism).
Both paradigms are designed to minimize further risk, prevent progressive functional
loss, and possibly reverse neurological deterioration. Clearly, in the case of an open
NTD, reversal of paralysis or sacral dysfunction is not expected or attained. Novel
repair strategies should be aimed at restoration of neurological function.
© 2005 by CRC Press LLC
10.5.1 FETAL SURGERY
Recent evidence suggests that the neurological deterioration associated with open
NTDs may have resulted from progressive intrauterine injury alone or in concert
with the primary defect of neurulation. For example, fetal ultrasonography revealed
that human fetuses with myelomeningoceles retained lower extremity movements
early in gestation and that the movements were lost by term.93 These data and
maternal reports that describe losses of fetal movements suggest that an event
occurring during gestation damaged fetal function.94
In the event of intrauterine injury, intrauterine intervention such as a surgical
repair may protect against progressive neurological deterioration. Animal models
designed with spina bifida were tested after intrauterine repair. Neurological function
was preserved in repaired animals.95 This result led to intrauterine repairs of open,
exposed spinal cords in humans.96,97
To determine the outcomes of fetal myelomeningocele repairs, the National
Institute of Child Health and Human Development (NICHD) sponsored the Management of Myelomeningocele Study (MOMS), a continuing clinical trial
[http://www.nichd.nih.gov]. Parameters undergoing study include optimal timing,
neurological recovery, and effects of repairs on associated hydrocephalus and Chiari
II malformations. The study is comparing two approaches to the treatment of babies
with spina bifida: surgery before birth (prenatal surgery) and the standard closure
surgery after birth (postnatal surgery). Preliminary results of human surgery show
failure to preserve fetal neurological function. Furthermore, when it appeared that
spinal cord function was present to a degree, it was less than predicted based on
data from the animal models.98 Improvements in the degree of hindbrain herniation
noted in the associated Chiari II malformation have also been demonstrated.99 Additionally, a reduction in the need for CSF shunting for hydrocephalus has been
Reported complications of fetal myelomeningocele surgery have been few; the
most common complication is preterm delivery.94 Major complications of intrauterine intervention such as maternal death from uterine rupture have been reported for
other types of fetal surgery.101 No uterine rupture resulting in maternal or fetal demise
has been reported to date for fetal myelomeningocele repair.94,97 Technical advancements, such as less invasive endoscopic procedures, have been proposed to avert
this severe complication.102,103
One key to predicting optimal outcomes of novel fetal surgery treatments for
myelomeningocele is understanding the structure of the placode. If the placode
retains normal patterning and is simply un-neurulated, a repair may be effective in
preventing secondary injury. There are mixed reports on whether placodes are normal
in animal models.104,105 Similar controversies surround human studies. Meuli et al.
characterized the human placode as having partial loss of tissue, containing hemorrhages and abrasions, while preserving developed elements of dorsal and ventral
parts of the spinal cord with nerve roots and ganglia.106 The abnormalities were
attributed to intrauterine injury.
Conversely, George and Cummings characterized the placode as having abnormal patterning along the dorsoventral and rostrocaudal axes indicative of a change
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in pattern determination and a paucity of maturing neurons with evidence of significant inflammatory infiltrate, gliosis, and fibrosis consistent with secondary injury.83
These data suggest that the myelomeningocele placode shows abnormal development
along with evidence of injury.
Reexamination of the animal model is needed to help clarify this controversy.
George and Fuh made several observations in a review.107 Two definitions of NTDs
were used to describe the surgical models: spina bifida or spina bifida-like and
surgical NTDs. All mammals except mice had spina bifida lesions in which the skin,
muscle, lamina, and dura were opened, but the spinal cord itself was not disturbed.108–112 Surgical NTDs were developed in avian species and mice; the dorsal
elements of the spinal cord were opened and splayed apart, and exposed the central
elements of the spinal cord to the surrounding environment.113–118 The surgical
models uncovered three mechanisms of injury:
1. Toxicity of the amniotic fluid
2. Direct intrauterine trauma
3. Developmental and growth distortion from laminectomy defect
Timing of lesions was critical. Spontaneous healing resulted if lesions occurred
early in gestation instead of later.111,114,117 Subsequent functional outcomes were
virtually indistinguishable between groups lesioned early in gestation and spontaneously healed and repaired fetuses lesioned later in gestation.83 Last, the surgical
animal models used were not the products of abnormal primary neurulation, and
could not directly address questions concerning the placode. These surgical models
represent a reopening mechanism of a closed neural tube that has not been shown
to appear in humans, but was reported in curtailed mouse mutants.119
The future of fetal surgery may rest in uncovering the mechanisms of fetal healing
and directly reconstituting the spinal cord. In the study of fetal wounds, healing was
demonstrated to occur rapidly and without scarring. The exact mechanisms of fetal
scarless healing remain unknown. However, transforming growth factor-beta and
hyaluronic acid-rich wound matrix play pivotal roles in scarless repair.120
The mechanism of annealing or healing that can lead to protection of the neural
tube has also not been defined. The fusion of reapproximated dorsal neural elements
in chicks has been suggested.118 A preliminary study in our laboratory utilizing
surgical NTDs in chicks and adding inhibitors of primary neurulation failed to
prevent reclosure of the neural tube (unpublished data). Therefore, reclosure in
chicks does not appear to be a recapitulation of primary neurulation. The underlying
molecular and cellular mechanisms that regulated the repair remain unclear, but the
ability of spinal cord cells to proliferate appeared important.118 These data suggest
that fetal interventions should be targeted at reinstituting mechanisms of fetal healing
that were turned off after a critical developmental phase.
10.5.2 SPINAL CORD REGENERATION
Current work on restoration of spinal cord function has focused on regeneration
after a spinal cord injury. If the precept from the fetal surgery is true, that the
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neurological sequelae in open NTDs are caused by intrauterine injuries, restoration
of cord function should be attainable. In fact, the majority of research has revealed
that an injured spinal cord can be restored by reconstituting or reestablishing molecular or cellular developmental mechanisms.121 Therefore, the developing spinal cord
appears to be the ideal substrate for regeneration of specific cell types and functional
connections as long as the milieu can be properly manipulated.
Paramount for the regeneration of the spinal cord is that the neuron becomes
“regeneration-capable” — it can restore the ability to demonstrate axonal growth
and proper targeting. A number of genes have been shown to be constitutively
expressed or upregulated in response to axonal growth. They have been termed
“regeneration-associated genes” and their products include transcription factors such
as c-jun, cytoskeleton components such as alpha tubulin, cytoplasmic growth cone
proteins such as GAP-43 and CAP-23, and cell adhesion molecules such as NCAM
and L1 that are important for growth cone guidance.122
The rate-limiting factor impacting regeneration is the inhibitory environment of
the mature CNS. CNS inhibition to axonal growth is broadly divided into nonpermissive factors related to myelin and the inhibitory nature of the gliotic scar. Proteins
identified in CNS myelin (NI-35 and NI-250) have been shown to function as neurite
inhibitory factors.123 At the injury site, dead cells, inflammation, and degraded tissue
are present. They contain reactive astrocytes, microglia, oligodendrocytes, and
meningeal cells that form gliotic scars that function as three-dimensional barriers to
As noted earlier, George and Cummings demonstrated that the myelomeningocele placode may have abnormal patterning along the dorsoventral and rostrocaudal
axes. This finding is indicative of a change in pattern determination, along with a
paucity of maturing neurons with evidence of significant inflammatory infiltrate,
gliosis, and fibrosis consistent with secondary injury.82 The impact that aberrant
development plays on the ability of the injured placode to regenerate and overcome
the inhibitory environment is unclear and remains a goal of future research.
Regenerative strategies in spinal cord injury include administration of trophic
factors, gene therapy, and cell transplantation. Intrathecal administration of trophic
factors such as neurotropin, nerve growth factor and glial-derived neurotrophic factor
upregulated growth cone proteins such as GAP-43 and CAP-23, propagated axonal
regrowth across an area of crush injury, and established functional connections.125
Interestingly, the administration of folate has been reported to assist in regenerating
axons in a spinal cord injury model via intraperitoneal administration (personal
communication). The mechanism of folate-assisted regeneration remains unknown.
Gene therapy strategies provide a way for longer lasting delivery of important
trophic factors. Trophic genes can be supplied ex vivo to an injured spinal cord by
inserting genetically altered cells that produce trophic factors.126 Another method is
applied in vivo: the neurotrophic gene is tranfected into the native spinal cord, usually
via a viral vector.127 Trophic factors listed above also serve as candidates for gene
therapy. Other classes of gene candidates are endogenous receptors or morphogens
important in embryonic development. For example, retinoic acid (RA) is important
in embryonic neural development69 and has been shown to stimulate embryonic
neurite outgrowth.128 RA administration failed to induce neurite growth in an injured
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adult spinal cord, presumably due to the lack of retinoic acid receptor-beta 2 (RAR
β2) upregulation.129 However, when the RAR β2 is upregulated, neurite outgrowth
can occur.130 Transfection into the adult spinal cord of RAR β2 alone was shown to
stimulate neurite outgrowth.130 Therefore, reinstitution of developmental mechanisms may be another methodology of cord regeneration.
Cellular transplantation strategies are aimed at circumventing the inhibitory
surround created by the gliotic scar. Candidates for transplantation are neural stem
cells and fetal cells that have the potential to develop into mature neurons or glia
and restore function by replacing or repairing axons and synaptic relays.131 Mature
cells such as Schwann cells or olfactory ensheathing cells also provide neurotrophic
support and myelination, thereby enhancing the regenerative environment. How the
myelomeningocele placode would respond to cellular transplantation remains
unclear. The lack of understanding of cell connectivity and patterning and the way
that environment responds to injury makes outcomes unpredictable, but unveils a
focal point for future study.
A final challenge to spinal cord regeneration of a NTD is that most of the studies
examined models of acute injury. Spinal cord dysfunction secondary in the congenital
setting is more likely to be chronic in nature. An important consideration in studies
of chronic injury is the survival of the injured neurons. Reports indicate that 25 to
50% of neurons die as early as 4 weeks postaxotomy,132 while the remaining cells
become atrophic. There is some evidence that trophic factors133 and fetal cell
transplants134 can enhance survival, even if applied 1 year after injury. Since many
patients with open and closed defects will present with neurological dysfunction
within this time frame, attempts at spinal cord regeneration remain viable techniques
10.6 SHOULD WE BELIEVE IN MAGIC?
The short answer is “yes.” Recent advances in genomics, proteomics, developmental
cell biology, biochemistry, embryology, neurobiology and neuroimaging have created the potential for a “golden age” in the cure of NTDs. Until then, NTDs remain
physically debilitating and are socioeconomic burdens. The time to advance neurosurgical management from supportive to restorative is now. It will be like magic!
1. Hrdlicka, A., Trephination among prehistoric people especially in America, Ciba
Foundation Symposium, 1:170–177, 1939.
2. Horsley, V., Brain surgery, Br. Med. J., 2:670–675, 1886.
3. Cushing, H., Intracranial tumors of preadolescence, Am. J. Dis. Children, 33:551–584,
4. Fulton, J., Harvey Cushing, Oxford, Blackwell Scientific, 1946.
5. Winston, K., Pediatric neurosurgery: pride and prejudice, Ped. Neurosurg., 32:58–68,
6. Begeer, I.H., Staal-Schreinemachers AL: the benefits of team treatment and control
of adult patients with spinal dysraphism, Eur. J. Ped. Surg., 6:15–16, 1996.
© 2005 by CRC Press LLC
7. Bowman, R.M., McLone D.G., Grant, J.A., et al., Spina bifida outcome: a 25-year
prospective, Ped. Neurosurg., 34:114–120, 2001.
8. Centers for Disease Control, Spina bifida incidence at birth: USA, 1983–1990,
WMMR, 41:497–500, 1992.
9. Reigel, D. and McLone, D.G., The tethered spinal cord, in Pediatric Neurosurgery,
Philadelphia, W.B. Saunders, 1994.
10. Copp, A., Brook, F.A., Estibeiro, J.P., Shum, A.S.W., and Cockroft, D.L., The embryonic development of mammalian NTDs, Prog. Neurobiol., 35:363–403, 1990.
11. Centers for Disease Control, Recommendations for the use of folic acid to reduce
the number of cases of spina bifida and other NTDs, WMMR, 41:1–7, 1992.
12. Holmes, L., Harris J., Oakley G.P., Jr. et al., Teratology Society consensus statement
on use of folic acid to reduce the risk of birth defects, Teratology, 55:381, 1997.
13. Milunsky, A., Jick, H., Jick, S. et al., Multivitamin/folic acid supplementation in early
pregnancy reduces the prevalence of NTDs, JAMA, 262:2847–2852, 1991.
14. Armstrong, D.C., Congenital malformations of the spine, Topics in Mag. Res. Imaging, 5:131–140, 1993.
15. Kirke, P.N., Mills, J.L., Whitehead, A.S. et al., Methylenetetrahydrofolate reductase
mutation and NTDs, Lancet, 348:1037–1038, 1996.
16. Van der Put, N.M., Gabreels, F., Stevens, E.M. et al., A second common mutation in
the methylenetetrahydrofolate reductase gene: an additional risk factor for neural tube
defects? [comment], Am. J. Human Gen., 62:1044–1051, 1998.
17. Speer, M.C., Worley, G., Mackey, J.F. et al., The thermolabile variant of methylenetetrahydrofolate reductase (MTHFR) is not a major risk factor for neural tube defect
in American Caucasians: NTD Collaborative Group, Neurogenetics, 1:149–150, 1997.
18. Anderson, R., Caveolae: where incoming and outgoing messengers meet, PNAS,
19. Waterland, R.A. and Jirtle, R.L., Transposable elements: targets for early nutritional
effects on epigenetic gene regulation, Mol. Cell. Biol., 23:5293–5300, 2003.
20. Bakker, R.C. and Brandjes, D.P., Hyperhomocysteinaemia and associated disease,
Pharm. World Sci., 19:126–132, 1997.
21. Blom, H.J., Mutated 5,10-methylenetetrahydrofolate reductase and moderate hyperhomocysteinaemia, Eur. J. Ped.,157:S131–134, 1998.
22. Burgoon, J.M., Selhub, J., Nadeau, M. et al., Investigation of the effects of folate
deficiency on embryonic development through the establishment of a folate deficient
mouse model, Teratology, 65:219–227, 2002.
23. Chatkupt, S., Chatkupt, S., and Johnson, W., Waardenburg syndrome and myelomeningocele in a family, J. Med. Gen., 30:83–84, 1993.
24. Epeldegui, M., Pena-Melian, A., Varela-Moreiras, G. et al., Homocysteine modifies
development of neurulation and dorsal root ganglia in chick embryos, Teratology,
25. Gos, M., Jr. and Szpecht-Potocka, A., Genetic basis of NTDs II: genes correlated
with folate and methionine metabolism, J. Appl. Gen., 43:511–524, 2002.
26. Castilla, E.E. and Orioli, I.M., Epidemiology of NTDs in South America, Am. J. Med.
27. Elwood, J., Little, J., and Elwood, J., Eds., Epidemiology and Control of NTDs,
Oxford, Oxford University Press, 1992.
28. Friel, J.K., Frecker, M., and Fraser, F.C., Nutritional patterns of mothers of children
with NTDs in Newfoundland, Am. J. Med. Gen., 55:195–199, 1995.
29. Jorde, L.B., Fineman, R.M., and Martin, R.A., Epidemiology of NTDs in Utah,
1940–1979, Am. J. Epidemiol., 119:487–495, 1984.
© 2005 by CRC Press LLC
30. Koch, M. and Fuhrmann, W., Epidemiology of NTDs in Germany, Hum. Gen.,
31. McDonnell, R.J., Johnson, Z., Delaney, V. et al., East Ireland, 1980–1994: epidemiology of NTDs, J. Epidemiol. Commun. Health, 53:782–788, 1999.
32. Xiao, K.A., Epidemiology of NTDs in China, Chung-Hua i Hsueh Tsa Chih (Chin.
Med. J.,) 69:189–191, 114, 1989.
33. Main, D. and Mennuti, M., NTDs: Issues in prenatal diagnosis and counseling,
Obstetr. Gynecol., 67:1–16, 1986.
34. Seller, M., Risk in spina bifida, Dev. Med. Child. Neurol., 36:1021–1025, 1994.
35. Seller, M.J., Vitamins, folic acid and the cause and prevention of NTDs, Ciba Found.
Symp., 181:161–173, 1994.
36. Lammer, E., Sever, L., and Oakley, G., Teratogen update: valproic acid, Teratology,
37. Robert, E. and Guibaud, P., Maternal valproic acid and congenital NTDs, Lancet,
38. Rosa, F., Spina bifida in infants of women treated with carbamazepine during pregnancy. New Engl. J. Med., 324:674–677, 1991.
39. Shaw, G., Velie, E., and Schaffer, D., Risk of neural tube defect-affected pregnancies
among obese women, JAMA, 275, 1996.
40. Watkins, M., Scanlon, K., Mulinare, J. et al., Is maternal obesity a risk factor for
anencephaly and spina bifida? Epidemiology, 7, 507–512, 1996.
41. Werler, M., Louik, C., Shapiro, S. et al., Prepregnant weight in relation to risk of
neural tube defects, JAMA, 275:1089–1092, 1996.
42. Harris, M. and Juriloff, D., Genetic landmarks for defects in mouse neural tube
closure, Teratology, 56:177–187, 1997.
43. Epstein, D.J., Malo, D., Vekemans, M. et al., Molecular characterization of a deletion
encompassing the splotch mutation on mouse chromosome 1, Genomics, 10:89–93,
44. Epstein, D.J., Vekemans, M., and Gros, P., Splotch (Sp2H), a mutation affecting
development of the mouse neural tube, shows a deletion within the paired homeodomain of Pax-3, Cell, 67:767–774, 1991.
45. Epstein, D.J., Vogan, K.J., Trasler, D.G. et al., A mutation within intron 3 of the Pax3 gene produces aberrantly spliced mRNA transcripts in the splotch (Sp) mouse
mutant, PNAS, 90:532–536, 1993.
46. Schimmang, T., Lemaistre, M., Vortkamp, A. et al., Expression of the zinc finger
gene Gli3 is affected in the morphogenetic mouse mutant extra-toes (Xt), Development, 116:799–804, 1992.
47. Wilson, V., Rashbass, P., and Beddington, R., Chimeric analysis of T (brachyury)
gene function, Development, 117:1321–1331, 1993.
48. Morrison-Graham, K., Schatteman, G., Bork, T. et al., A PDGF receptor mutation in
the mouse (patch) perturbs the development of a non-neuronal subset of neural crestderived cells, Development, 115:133–142, 1992.
49. Homanics, G., Smith, T., Zhang, S. et al., Targeted modification of the apolipoprotein
B gene results in hypobetalipoproteinemia and developmental abnormalities in mice,
PNAS, 90:2389–2393, 1993.
50. Lufkin, T., Dierich, A., LeMeur, M. et al., Disruption of the Hox-1.6 homeobox gene
results in defects in a region corresponding to its rostral domain of expression, Cell,
51. Moase, C.E. and Trasler, D.G., Splotch locus mouse mutants: models for NTDs and
Waardenburg syndrome type I in humans, J. Med. Gen., 29:145–151, 1992.
© 2005 by CRC Press LLC
52. Hui, C. and Joyner, A., A mouse of Greig cephalopolysyndactyly syndrome: the extra
toes mutation contains an intragenic delection of the Gli3 gene, Nature Gen.,
53. Morrison, K., Papapetrou, C., Attwood, J. et al., Genetic mapping of the human
homologue (T) of mouse T (brachyury) and a search for allele association between
human T and spina bifida, Hum. Mol. Gen., 5:669–674, 1996.
54. Melvin, E.C., George, T.M., Worley, G. et al., Genetic studies in NTDs: NTD Collaborative Group, Ped. Neurosurg., 32:1–9, 2000.
55. Yamada, T., Caudalization by the amphibian organizer: brachyury, convergent extension and retinoic acid, Development, 120:3051–3062, 1994.
56. Cooke, J., Takada, S., and McMahon, A., Experimental control of axial pattern in
the chick bastoderm by local expression of Wnt and activin: role of HNK-1-positive
cells, Dev. Biol., 164:513–527, 1994.
57. Jacobson, A., Inductive processes in embryonic development, Science, 152:25–34,
58. Jessell, T. and Melton, D., Diffusible factors in vertebrate embryonic induction, Cell,
59. Kispert, A., Ortner, H., Cooke, J., and Herrmann, B.G., The chick brachyury gene:
developmental expression pattern and response to axial induction by localized activin,
Dev. Biol., 168:406–415, 1995.
60. Knezevic, V., Ranson, M., and Mackem, S., The organizer-associated chick homeobox
gene, Gnot1, is expressed before gastrulation and regulated synergistically by activin
and retinoic acid, Dev. Biol., 171:458–470, 1995.
61. Menkes, B. and Sandor, S., Researches on the development of axial organs, Roum
Rev. Embryol. Cytol., 6:65–88, 1969.
62. Mitrani, E. and Shimoni, Y., Retinoic acid inhibits growth in agarose of early chick
embryonic cells and may be involved in regulation of axis formation, Development,
63. Mitrani, E.G.Y., Shohat, H., and Ziv, T., Fibroblast growth factor during mesoderm
induction in the early chick embryo, Development, 109:387–393, 1990.
64. Ziv, T., Shimoni, Y., and Mitrani, E., Activin can generate ectopic axial structures in
chick blastoderm explants, Development, 115:689–694, 1992.
65. Belting, H.G., Hauptmann, G., Meyer, D. et al., Spiel ohne grenzen/pou2 is required
during establishment of the zebrafish midbrain-hindbrain boundary organizer, Development, 128:4165–4176, 2001.
66. Deardorff, M.A., Tan, C., Conrad, L.J., et al., Frizzled-8 is expressed in the Spemann
organizer and plays a role in early morphogenesis, Development, 125:2687–2700,
67. Zhao, Q., Behringer, R.R., and de Crombrugghe, B., Prenatal folic acid treatment
suppresses acrania and meroanencephaly in mice mutant for the Cart1 homeobox
gene, Nature Gen., 13:275–283, 1996.
68. Zinyk, D.L., Mercer, E.H., Harris, E. et al., Fate mapping of the mouse midbrain–hindbrain constriction using a site-specific recombination system, Curr. Biol., 8:665–668,
69. Maden, M. and Holder, N., The involvement of retinoic acid in the development of
the vertebrate central nervous system, Dev. Suppl., 2:87–94, 1991.
70. Yamada, T., Pfaff, S.L., Edlund, T., and Jessell, T.M., Control of cell pattern in the
neural tube: motor neuron induction by diffusible factors from notochord and floor
plate, Cell, 73:673–686, 1993.
© 2005 by CRC Press LLC
71. Schoenwolf, G.C. and Smith, J.L., Mechanisms of neurulation: traditional viewpoint
and recent advances, Development,109:243–270, 1990.
72. Griffith, C.M., Wiley, M.J., and Sanders, E.J., The vertebrate tail bud: three germ
layers from one tissue, Anat. Embryol., 185:101–113, 1992.
73. Muller, F. and O’Rahilly, R., The development of the human brain, the closure of the
caudal neuropore, and the beginning of secondary neurulation at stage 12, Anat.
Embryol., 176:413–430, 1987.
74. Nievelstein, R.A., Hartwig, N.G., Vermeij-Keers, C. et al., Embryonic development
of the mammalian caudal neural tube, Teratology, 48:21–31, 1993.
75. O’Rahilly, R. and Muller, F., Neurulation in the normal human embryo, Ciba Found.
Symp., 181:70–82, 1994.
76. Schoenwolf, G.C., Histological and ultrastructural studies of secondary neurulation
in mouse embryos, Am. J. Anat., 169:361–376, 1984.
77. Schoenwolf, G.C. and Delongo, J., Ultrastructure of secondary neurulation in the
chick embryo, American J. Anat., 158:43–63, 1980.
78. Yang, H.J., Wang, K.C., Chi, J.G. et al., Neural differentiation of caudal cell mass
(secondary neurulation) in chick embryos: Hamburger and Hamilton Stages 16–45,
Brain Res. Dev. Brain Res., 142:31–36, 2003.
79. Catala, M., Teillet, M.A., De Robertis, E.M. et al., A spinal cord fate map in the
avian embryo: while regressing, Hensen’s node lays down the notochord and floor
plate thus joining the spinal cord lateral walls, Development, 122:2599–2610, 1996.
80. Le Douarin, N.M., Teillet, M.A., and Catala, M., Neurulation in amniote vertebrates:
a novel view deduced from the use of quail-chick chimeras, Int. J. Dev. Biol.,
81. Cummings, T. and George, T., The immunohistochemical profile of the normal conus
medullaris and the filum terminale, Neuroembryology (in press), 2003.
82. George, T.M., Bulsara, K.R., and Cummings, T.J., The immunohistochemical profile
of the tethered filum terminale, Ped. Neurosurg., 39:227–233, 2003.
83. George, T.M. and Cummings, T.J., Immunohistochemical profile of the myelomeningocele placode: is the placode normal? Ped. Neurosurg., 39:234–239, 2003.
84. Carter, M., Ulrich, S., Oofuji, Y. et al., Crooked tail (Cd) models human folateresponsive NTDs, Hum. Mol. Gen., 8:2199–2204, 1999.
85. Fleming, A. and Copp, A.J., Embryonic folate metabolism and mouse NTDs, Science,
86. Kowal, M. and Wojcierowski, J., Role and significance of DNA methylation, Polski
Merkur/ Lekarsk., 14:364–368, 2003.
87. Reik, W., Dean, W., and Walter, J., Epigenetic reprogramming in mammalian development, Science, 293:1089–1093, 2001.
88. Van den Veyver, I.B., Genetic effects of methylation diets, Annu. Rev. Nutr.,
89. Rakyan, V.K., Blewitt, M.D., Druker, R. et al., Metastable epialleles in mammals,
Trends Gen., 18:348–351, 2002.
90. Cavalli, P. and Copp, A.J., Inositol and folate-resistant NTDs, J. Med. Gen., 39:E5,
91. Greene, N.D. and Copp, A.J., Inositol prevents folate-resistant NTDs in the mouse,
Nature Med., 3:60–66, 1997.
92. Corcoran, J., What are the molecular mechanisms of NTDs? Bioessays, 20:6–8, 1998.
93. Korenromp, M.J., van Gool, J.D., Bruinese, H.W. et al., Early fetal leg movements
in myelomeningocele, Lancet, 1:917–918, 1986.
© 2005 by CRC Press LLC