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5 Post-transcriptional Control: The Role of miRNAs
F. Cavodeassi et al.
expression of downstream genes and the correct establishment of axis polarity (Kim
et al. 2011). An additional example of how miRNAs impact on AP patterning is
provided by their role in reﬁning and maintaining the function of secondary
organisers, such as in the case of the MHB. miR-9 is expressed around this organizing boundary, where it targets several transducers of the FGF signalling pathway
(Leucht et al. 2008), limiting its signalling effects. As miRNAs usually bind several
distinct mRNAs, miR-9 seems also to reduce the activity of neurogenic genes at the
MHB, maintaining this territory in an undifferentiated state essential for its function
as an organiser (Leucht et al. 2008).
Quite likely miRNAs participate in the regionalization of the entire forebrain
primordium, but at the moment, most studies have focused on the eye primordium,
which expresses several miRNAs and is severely affected by Dicer inactivation.
Dicer deletion causes microphthalmia (reduction of the eye size) affecting the lens
placode, the neural retina, the pathﬁnding of the retinal ganglion cell axons as well
as the pigmentation and adhesion of the Retinal Pigment Epithelium (RPE), which,
in turn, affect photoreceptors’ maturation (Conte et al. 2013; Ohana et al. 2015).
Besides the general demonstration that mRNA silencing is relevant for eye speciﬁcation, knock-down/out studies are beginning to delineate the speciﬁc function of
each miRNA in eye formation. Among them, miR-124 and miR-204 are particularly important. miR-124 maintains optic vesicle cell proliferation at early stages of
development by turning off the proneural gene neuroD1. This early function prevents the onset of neurogenesis (Liu et al. 2011). Later on miR-124 promotes
differentiated cone photoreceptor survival by targeting the TF Lhx2 (Sanuki et al.
2011). miR-204 instead modulates the levels of the TF Meis2, which is upstream of
Pax6 in the GRNs controlling morphogenesis and speciﬁcation of both the lens and
the retina (Conte et al. 2010). Consistent with the general observation that miRNAs
have rather heterogeneous targets, slightly later, miR-204 targets EphB2 and EfnB3
(Conte et al. 2014), a signalling system implicated in retinal ganglion cell axon
pathﬁnding, as well as effector genes of RPE differentiation (Adijanto et al. 2012).
Many more studies are needed to fully understand how miRNAs contribute to
forebrain development. Nevertheless, it is becoming apparent that many miRNAs
can contribute to the regulation of the same process and also that each miRNA, is
recurrently used during development for different purposes, further contributing to
diversify the GRNs that lead to a mature forebrain.
In conclusion, in this chapter we have provided a general and simpliﬁed view of the
principles that govern early forebrain development. This information derives from a
huge number of studies based on experimental manipulations of gene activity in
different vertebrate species, of which unfortunately we could not give a full account
here. These studies have been facilitated by the sequencing of several genomes,
which also led to the identiﬁcation of a large number of non-coding RNAs, as well
Principles of Early Vertebrate Forebrain Formation
as of the presence of evolutionary conserved non-coding regions. The latter ﬁnding,
in turn, has uncovered the existence of highly speciﬁc regulatory codes that deﬁne
the dynamic expression of key developmental genes, enabling the assembly GRN
models (reviewed in Nord et al. 2015). These models together with technical
advances in embryonic imaging have been of enormous value to couple gene
activity to the dynamics of forebrain development.
From this manifold experimental work a few general principles have emerged.
That forebrain development occurs in a rather parsimonious way is likely the most
evident of these principles. Indeed, a reduced set of genes is constantly repurposed
to obtain different outcomes in different regions of the forebrain either through
combinations with different network partners, interactions with different co-factors
or variations in exposure to and amount of gene product. A second important
principle is that “kernel” components of the forebrain GRNs are extremely conserved across evolution and their inactivation result in profound alterations or loss
of the forebrain primordium. Effector genes instead are less constrained and have
undergone variations especially in their regulatory regions, which is thought to have
favoured the progressive evolution of the vertebrate forebrain. Of particular relevance, recent studies have shown that human regulatory elements exhibit high
levels of evolutionary innovation both in sequence and function (reviewed in Nord
et al. 2015).
An additional important aspect underlying progressive forebrain development is
the contribution of cytoskeletal rearrangements and of the evolving cell interactions,
which both couple patterning and morphogenesis. These contributions are still
poorly understood but their elucidation should give hints on how different vertebrate species have adopted distinct cell arrangements to reach the same ﬁnal result.
The formation of the neural tube or of the eye in mouse/chick and teleost ﬁshes are
example of these differences.
Despite these rather impressive advances, much still needs to be understood
towards a full comprehension of how the forebrain forms. The array of genes
involved is likely incomplete and the assembly of the GRNs is still rudimentary
(Nord et al. 2015). How the different effectors of the GRN contribute to neuroepithelial cell patterning and sorting need much attention. For example, we have
gained knowledge on the importance of adhesive mechanisms but we know little on
how adhesive events are interrupted and virtually nothing on the possible contribution that the so called house-keeping functions might have on the acquisition of
forebrain cell identity. Is metabolism, energy production or even response to
external stimuli relevant to forebrain morphogenesis? An intriguing study has
shown that light perceived in utero influences eye developmental events (Rao et al.
2013), making these questions worthwhile to be addressed.
An important aspect is how much of what we learn from organisms can be
applied to human forebrain development. The outstanding advances in the use of
ES and iPSC cells to reproduce organ formation in culture offer an important tool to
answer such a question. For example, comparative analysis of mouse and human
eye organoids has shown intrinsic differences of the assembled eyes according to
the respective species (Eiraku et al. 2011; Nakano et al. 2012), including the
F. Cavodeassi et al.
generation of a proportion of cone or rod photoreceptors, according to the
respective nocturnal and diurnal type of vision of mice and humans.
As many tools are now in place, we should expect a rapid broadening of our
knowledge on forebrain development that will help to decipher the causes of the
many still poorly understood pathologies linked to congenital alterations of the
Acknowledgments Work in our lab is supported by grants from the Spanish Government
MINECO (BFU2014-55918-P to F.C.; BFU-2013-43213-P and BFU2014-55738-REDT to P.B.),
the European Commission (CIG321788 to F.C. and P.B.); the Comunidad Autonoma de Madrid
(CAM; S2010/BMD-2315 to P.B.); the CIBERER, ISCIII to P.B. and by an Institutional Grant
from the Fundación Ramon Areces.
Acampora, D., Mazan, S., Lallemand, Y., Avantaggiato, V., Maury, M., Simeone, A., et al. (1995).
Forebrain and midbrain regions are deleted in Otx2−/− mutants due to a defective anterior
neuroectoderm speciﬁcation during gastrulation. Development, 121, 3279–3290.
Adijanto, J., Castorino, J. J., Wang, Z. X., Maminishkis, A., Grunwald, G. B., & Philp, N. J. (2012).
Microphthalmia-associated transcription factor (MITF) promotes differentiation of human
retinal pigment epithelium (RPE) by regulating microRNAs-204/211 expression. The Journal of
Biological Chemistry, 287, 20491–20503.
Andoniadou, C. L., & Martinez-Barbera, J. P. (2013). Developmental mechanisms directing early
anterior forebrain speciﬁcation in vertebrates. Cellular and Molecular Life Sciences: CMLS,
Araya, C., Tawk, M., Girdler, G. C., Costa, M., Carmona-Fontaine, C., & Clarke, J. D. (2014).
Mesoderm is required for coordinated cell movements within zebraﬁsh neural plate in vivo.
Neural Development, 9, 9.
Beccari, L., Conte, I., Cisneros, E., & Bovolenta, P. (2012). Sox2-mediated differential activation
of Six3.2 contributes to forebrain patterning. Development, 139, 151–164.
Beccari, L., Marco-Ferreres, R., & Bovolenta, P. (2013). The logic of gene regulatory networks in
early vertebrate forebrain patterning. Mechanisms of Development, 130, 95–111.
Beccari, L., Marco-Ferreres, R., Tabanera, N., Manfredi, A., Souren, M., Wittbrodt, B., et al.
(2015). A trans-regulatory code for the forebrain expression of Six3.2 in the medaka ﬁsh. The
Journal of Biological Chemistry.
Bernstein, E., Kim, S. Y., Carmell, M. A., Murchison, E. P., Alcorn, H., Li, M. Z., et al. (2003).
Dicer is essential for mouse development. Nature Genetics, 35, 215–217.
Bhinge, A., Poschmann, J., Namboori, S. C., Tian, X., Jia Hui Loh, S., Traczyk, A., et al. (2014).
MiR-135b is a direct PAX6 target and speciﬁes human neuroectoderm by inhibiting
TGF-beta/BMP signaling. The EMBO Journal, 33, 1271–1283.
Bielen, H., & Houart, C. (2012). BMP signaling protects telencephalic fate by repressing eye
identity and its Cxcr4-dependent morphogenesis. Developmental Cell, 23, 812–822.
Blaess, S., Szabo, N., Haddad-Tovolli, R., Zhou, X., & Alvarez-Bolado, G. (2014). Sonic
hedgehog signaling in the development of the mouse hypothalamus. Frontiers in
Neuroanatomy, 8, 156.
Bovolenta, P., Esteve, P., Ruiz, J. M., Cisneros, E., & Lopez-Rios, J. (2008). Beyond Wnt
inhibition: New functions of secreted Frizzled-related proteins in development and disease.
Journal of Cell Science, 121, 737–746.
Principles of Early Vertebrate Forebrain Formation
Braun, M. M., Etheridge, A., Bernard, A., Robertson, C. P., & Roelink, H. (2003). Wnt signaling
is required at distinct stages of development for the induction of the posterior forebrain.
Development, 130, 5579–5587.
Brown, K. E., Keller, P. J., Ramialison, M., Rembold, M., Stelzer, E. H., Loosli, F., & Wittbrodt,
J. (2010). Nlcam modulates midline convergence during anterior neural plate morphogenesis.
Developmental Biology, 339, 14–25.
Buckley, C., & Clarke, J. (2014). Establishing the plane of symmetry for lumen formation and
bilateral brain formation in the zebraﬁsh neural rod. Seminars in Cell & Developmental
Biology, 31, 100–105.
Cavodeassi, F. (2014). Integration of anterior neural plate patterning and morphogenesis by the
Wnt signaling pathway. Developmental Neurobiology, 74, 759–771.
Cavodeassi, F., Carreira-Barbosa, F., Young, R. M., Concha, M. L., Allende, M. L., Houart, C.,
et al. (2005). Early stages of zebraﬁsh eye formation require the coordinated activity of Wnt11,
Fz5, and the Wnt/beta-catenin pathway. Neuron, 47, 43–56.
Cavodeassi, F., & Houart, C. (2012). Brain regionalization: Of signaling centers and boundaries.
Developmental Neurobiology, 72, 218–233.
Cavodeassi, F., Ivanovitch, K., & Wilson, S. W. (2013). Eph/Ephrin signalling maintains eye ﬁeld
segregation from adjacent neural plate territories during forebrain morphogenesis.
Development, 140, 4193–4202.
Ciruna, B., Jenny, A., Lee, D., Mlodzik, M., & Schier, A. F. (2006). Planar cell polarity signalling
couples cell division and morphogenesis during neurulation. Nature, 439, 220–224.
Clarke, J. (2009). Role of polarized cell divisions in zebraﬁsh neural tube formation. Current
Opinion in Neurobiology, 19, 134–138.
Conte, I., Banﬁ, S., & Bovolenta, P. (2013). Non-coding RNAs in the development of sensory
organs and related diseases. Cellular and Molecular Life Sciences: CMLS, 70, 4141–4155.
Conte, I., Carrella, S., Avellino, R., Karali, M., Marco-Ferreres, R., Bovolenta, P., et al. (2010).
miR-204 is required for lens and retinal development via Meis2 targeting. Proceedings of the
National Academy of Sciences USA, 107, 15491–15496.
Conte, I., Merella, S., Garcia-Manteiga, J. M., Migliore, C., Lazarevic, D., Carrella, S., et al.
(2014). The combination of transcriptomics and informatics identiﬁes pathways targeted by
miR-204 during neurogenesis and axon guidance. Nucleic Acids Research, 42, 7793–7806.
Diaz, N. F., Cruz-Resendiz, M. S., Flores-Herrera, H., Garcia-Lopez, G., & Molina-Hernandez, A.
(2014). MicroRNAs in central nervous system development. Reviews in the Neurosciences, 25,
Du, Z. W., Ma, L. X., Phillips, C., & Zhang, S. C. (2013). miR-200 and miR-96 families repress
neural induction from human embryonic stem cells. Development, 140, 2611–2618.
Eiraku, M., Takata, N., Ishibashi, H., Kawada, M., Sakakura, E., Okuda, S., et al. (2011).
Self-organizing optic-cup morphogenesis in three-dimensional culture. Nature, 472, 51–56.
Erwin, D. H., & Davidson, E. H. (2009). The evolution of hierarchical gene regulatory networks.
Nature Reviews Genetics, 10, 141–148.
Esteve, P., & Bovolenta, P. (2006). Secreted inducers in vertebrate eye development: More
functions for old morphogens. Current Opinion in Neurobiology, 16, 13–19.
Esteve, P., Lopez-Rios, J., & Bovolenta, P. (2004). SFRP1 is required for the proper establishment
of the eye ﬁeld in the medaka ﬁsh. Mechanisms of Development, 121, 687–701.
Esteve, P., Sandonis, A., Cardozo, M., Malapeira, J., Ibanez, C., Crespo, I., et al. (2011a). SFRPs
act as negative modulators of ADAM10 to regulate retinal neurogenesis. Nature Neuroscience,
Esteve, P., Sandonis, A., Ibanez, C., Shimono, A., Guerrero, I., & Bovolenta, P. (2011b). Secreted
frizzled-related proteins are required for Wnt/beta-catenin signalling activation in the vertebrate
optic cup. Development, 138, 4179–4184.
Giraldez, A. J., Cinalli, R. M., Glasner, M. E., Enright, A. J., Thomson, J. M., Baskerville, S., et al.
(2005). MicroRNAs regulate brain morphogenesis in zebraﬁsh. Science, 308, 833–838.
F. Cavodeassi et al.
Glinka, A., Wu, W., Delius, H., Monaghan, A. P., Blumenstock, C., & Niehrs, C. (1998).
Dickkopf-1 is a member of a new family of secreted proteins and functions in head induction.
Nature, 391, 357–362.
Hammond, S. M., Bernstein, E., Beach, D., & Hannon, G. J. (2000). An RNA-directed nuclease
mediates post-transcriptional gene silencing in Drosophila cells. Nature, 404, 293–296.
Hirata, T., Nakazawa, M., Muraoka, O., Nakayama, R., Suda, Y., & Hibi, M. (2006). Zinc-ﬁnger
genes Fez and Fez-like function in the establishment of diencephalon subdivisions.
Development, 133, 3993–4004.
Houart, C., Caneparo, L., Heisenberg, C., Barth, K., Take-Uchi, M., & Wilson, S. (2002).
Establishment of the telencephalon during gastrulation by local antagonism of Wnt signaling.
Neuron, 35, 255–265.
Hutvagner, G., McLachlan, J., Pasquinelli, A. E., Balint, E., Tuschl, T., & Zamore, P. D. (2001).
A cellular function for the RNA-interference enzyme Dicer in the maturation of the let-7 small
temporal RNA. Science, 293, 834–838.
Ivanovitch, K., Cavodeassi, F., & Wilson, S. W. (2013). Precocious acquisition of neuroepithelial
character in the eye ﬁeld underlies the onset of eye morphogenesis. Developmental Cell, 27,
Jeong, J. Y., Einhorn, Z., Mathur, P., Chen, L., Lee, S., Kawakami, K., et al. (2007). Patterning the
zebraﬁsh diencephalon by the conserved zinc-ﬁnger protein Fezl. Development, 134, 127–136.
Kapsimali, M., Kloosterman, W. P., de Bruijn, E., Rosa, F., Plasterk, R. H., & Wilson, S. W.
(2007). MicroRNAs show a wide diversity of expression proﬁles in the developing and mature
central nervous system. Genome Biology, 8, R173.
Kaspi, H., Chapnik, E., Levy, M., Beck, G., Hornstein, E., & Soen, Y. (2013). Brief report:
miR-290-295 regulate embryonic stem cell differentiation propensities by repressing Pax6.
Stem Cells, 31, 2266–2272.
Kawase-Koga, Y., Otaegi, G., & Sun, T. (2009). Different timings of Dicer deletion affect
neurogenesis and gliogenesis in the developing mouse central nervous system. Developmental
Dynamics: An Ofﬁcial Publication of the American Association of Anatomists, 238,
Kiecker, C., & Lumsden, A. (2012). The role of organizers in patterning the nervous system.
Annual Review of Neuroscience, 35, 347–367.
Kim, N. H., Kim, H. S., Kim, N. G., Lee, I., Choi, H. S., Li, X. Y., et al. (2011). p53 and
microRNA-34 are suppressors of canonical Wnt signaling. Science Signaling, 4, ra71.
Kobayashi, D., Kobayashi, M., Matsumoto, K., Ogura, T., Nakafuku, M., & Shimamura, K.
(2002). Early subdivisions in the neural plate deﬁne distinct competence for inductive signals.
Development, 129, 83–93.
Kobayashi, K., Luo, M., Zhang, Y., Wilkes, D. C., Ge, G., Grieskamp, T., et al. (2009). Secreted
Frizzled-related protein 2 is a procollagen C proteinase enhancer with a role in ﬁbrosis
associated with myocardial infarction. Nature Cell Biology, 11, 46–55.
Kudoh, T., Concha, M. L., Houart, C., Dawid, I. B., & Wilson, S. W. (2004). Combinatorial Fgf
and Bmp signalling patterns the gastrula ectoderm into prospective neural and epidermal
domains. Development, 131, 3581–3592.
Kutejova, E., Briscoe, J., & Kicheva, A. (2009). Temporal dynamics of patterning by morphogen
gradients. Current Opinion in Genetics & Development, 19, 315–322.
Lagutin, O. V., Zhu, C. C., Kobayashi, D., Topczewski, J., Shimamura, K., Puelles, L., et al.
(2003). Six3 repression of Wnt signaling in the anterior neuroectoderm is essential for
vertebrate forebrain development. Genes & Development, 17, 368–379.
Lee, H. X., Ambrosio, A. L., Reversade, B., & De Robertis, E. M. (2006). Embryonic dorsalventral signaling: Secreted frizzled-related proteins as inhibitors of tolloid proteinases. Cell,
Lee, Y., Ahn, C., Han, J., Choi, H., Kim, J., Yim, J., et al. (2003). The nuclear RNase III Drosha
initiates microRNA processing. Nature, 425, 415–419.
Lee, Y., Jeon, K., Lee, J. T., Kim, S., & Kim, V. N. (2002). MicroRNA maturation: Stepwise
processing and subcellular localization. The EMBO Journal, 21, 4663–4670.
Principles of Early Vertebrate Forebrain Formation
Leucht, C., Stigloher, C., Wizenmann, A., Klafke, R., Folchert, A., & Bally-Cuif, L. (2008).
MicroRNA-9 directs late organizer activity of the midbrain-hindbrain boundary. Nature
Neuroscience, 11, 641–648.
Li, J. Y., Lao, Z., & Joyner, A. L. (2005). New regulatory interactions and cellular responses in the
isthmic organizer region revealed by altering Gbx2 expression. Development, 132, 1971–1981.
Linker, C., & Stern, C. D. (2004). Neural induction requires BMP inhibition only as a late step,
and involves signals other than FGF and Wnt antagonists. Development, 131, 5671–5681.
Liu, K., Liu, Y., Mo, W., Qiu, R., Wang, X., Wu, J. Y., et al. (2011). MiR-124 regulates early
neurogenesis in the optic vesicle and forebrain, targeting NeuroD1. Nucleic Acids Research,
Liu, W., Lagutin, O., Swindell, E., Jamrich, M., & Oliver, G. (2010). Neuroretina speciﬁcation in
mouse embryos requires Six3-mediated suppression of Wnt8b in the anterior neural plate. The
Journal of Clinical Investigation, 120, 3568–3577.
Longabaugh, W. J., Davidson, E. H., & Bolouri, H. (2005). Computational representation of
developmental genetic regulatory networks. Developmental Biology, 283, 1–16.
Lopez-Rios, J., Esteve, P., Ruiz, J. M., & Bovolenta, P. (2008). The Netrin-related domain of
Sfrp1 interacts with Wnt ligands and antagonizes their activity in the anterior neural plate.
Neural Development, 3, 19.
Lowery, L. A., & Sive, H. (2004). Strategies of vertebrate neurulation and a re-evaluation of
teleost neural tube formation. Mechanisms of Development, 121, 1189–1197.
Makeyev, E. V., Zhang, J., Carrasco, M. A., & Maniatis, T. (2007). The MicroRNA miR-124
promotes neuronal differentiation by triggering brain-speciﬁc alternative pre-mRNA splicing.
Molecular Cell, 27, 435–448.
Marti, E., & Bovolenta, P. (2002). Sonic hedgehog in CNS development: One signal, multiple
outputs. Trends in Neurosciences, 25, 89–96.
Martinez-Barbera, J. P., Signore, M., Boyl, P. P., Puelles, E., Acampora, D., Gogoi, R., et al.
(2001). Regionalisation of anterior neuroectoderm and its competence in responding to
forebrain and midbrain inducing activities depend on mutual antagonism between OTX2 and
GBX2. Development, 128, 4789–4800.
Martinez-Morales, J. R. (2015). Toward understanding the evolution of vertebrate gene regulatory
networks: Comparative genomics and epigenomic approaches. Brieﬁngs in Functional
Matsumoto, K., Nishihara, S., Kamimura, M., Shiraishi, T., Otoguro, T., Uehara, M., et al. (2004).
The prepattern transcription factor Irx2, a target of the FGF8/MAP kinase cascade, is involved
in cerebellum formation. Nature Neuroscience, 7, 605–612.
Matsuo, I., Kuratani, S., Kimura, C., Takeda, N., & Aizawa, S. (1995). Mouse Otx2 functions in
the formation and patterning of rostral head. Genes & Development, 9, 2646–2658.
Maurus, D., Heligon, C., Burger-Schwarzler, A., Brandli, A. W., & Kuhl, M. (2005).
Noncanonical Wnt-4 signaling and EAF2 are required for eye development in Xenopus
laevis. The EMBO Journal, 24, 1181–1191.
Nakano, T., Ando, S., Takata, N., Kawada, M., Muguruma, K., Sekiguchi, K., et al. (2012).
Self-formation of optic cups and storable stratiﬁed neural retina from human ESCs. Cell Stem
Cell, 10, 771–785.
Nishimura, T., Honda, H., & Takeichi, M. (2012). Planar cell polarity links axes of spatial
dynamics in neural-tube closure. Cell, 149, 1084–1097.
Nord, A. S., Pattabiraman, K., Visel, A., & Rubenstein, J. L. (2015). Genomic perspectives of
transcriptional regulation in forebrain development. Neuron, 85, 27–47.
Ohana, R., Weiman-Kelman, B., Raviv, S., Tamm, E. R., Pasmanik-Chor, M., Rinon, A., et al.
(2015). MicroRNAs are essential for differentiation of the retinal pigmented epithelium and
maturation of adjacent photoreceptors. Development, 142, 2487–2498.
Okuda, Y., Ogura, E., Kondoh, H., & Kamachi, Y. (2010). B1 SOX coordinate cell speciﬁcation
with patterning and morphogenesis in the early zebraﬁsh embryo. PLoS Genetics, 6, e1000936.
Okuda, Y., Yoda, H., Uchikawa, M., Furutani-Seiki, M., Takeda, H., Kondoh, H., et al. (2006).
Comparative genomic and expression analysis of group B1 sox genes in zebraﬁsh indicates
F. Cavodeassi et al.
their diversiﬁcation during vertebrate evolution. Developmental Dynamics: An Ofﬁcial
Publication of the American Association of Anatomists, 235, 811–825.
Ozair, M. Z., Kintner, C., & Brivanlou, A. H. (2013). Neural induction and early patterning in
vertebrates. Wiley Interdisciplinary Reviews. Developmental Biology, 2, 479–498.
Pera, E. M., Ikeda, A., Eivers, E., & De Robertis, E. M. (2003). Integration of IGF, FGF, and
anti-BMP signals via Smad1 phosphorylation in neural induction. Genes & Development, 17,
Rao, S., Chun, C., Fan, J., Kofron, J. M., Yang, M. B., Hegde, R. S., et al. (2013). A direct and
melanopsin-dependent fetal light response regulates mouse eye development. Nature, 494,
Rembold, M., Loosli, F., Adams, R. J., & Wittbrodt, J. (2006). Individual cell migration serves as
the driving force for optic vesicle evagination. Science, 313, 1130–1134.
Rhinn, M., Lun, K., Ahrendt, R., Geffarth, M., & Brand, M. (2009). Zebraﬁsh gbx1 reﬁnes the
midbrain-hindbrain boundary border and mediates the Wnt8 posteriorization signal. Neural
Development, 4, 12.
Rodriguez-Seguel, E., Alarcon, P., & Gomez-Skarmeta, J. L. (2009). The Xenopus Irx genes are
essential for neural patterning and deﬁne the border between prethalamus and thalamus through
mutual antagonism with the anterior repressors Fezf and Arx. Developmental Biology, 329,
Rubenstein, J. L., Shimamura, K., Martinez, S., & Puelles, L. (1998). Regionalization of the
prosencephalic neural plate. Annual Review of Neuroscience, 21, 445–477.
Sanchez-Arrones, L., Ferran, J. L., Rodriguez-Gallardo, L., & Puelles, L. (2009). Incipient
forebrain boundaries traced by differential gene expression and fate mapping in the chick
neural plate. Developmental Biology, 335, 43–65.
Sanuki, R., Onishi, A., Koike, C., Muramatsu, R., Watanabe, S., Muranishi, Y., et al. (2011).
miR-124a is required for hippocampal axogenesis and retinal cone survival through Lhx2
suppression. Nature Neuroscience, 14, 1125–1134.
Sasai, Y., Eiraku, M., & Suga, H. (2012). In vitro organogenesis in three dimensions:
Self-organising stem cells. Development, 139, 4111–4121.
Scholpp, S., Foucher, I., Staudt, N., Peukert, D., Lumsden, A., & Houart, C. (2007). Otx1l, Otx2
and Irx1b establish and position the ZLI in the diencephalon. Development, 134, 3167–3176.
Schwarz, D. S., Hutvagner, G., Du, T., Xu, Z., Aronin, N., & Zamore, P. D. (2003). Asymmetry in
the assembly of the RNAi enzyme complex. Cell, 115, 199–208.
Shinozaki, K., Yoshida, M., Nakamura, M., Aizawa, S., & Suda, Y. (2004). Emx1 and Emx2
cooperate in initial phase of archipallium development. Mechanisms of Development, 121,
Sokol, S. Y. (2015). Spatial and temporal aspects of Wnt signaling and planar cell polarity during
vertebrate embryonic development. Seminars in Cell & Developmental Biology, 42, 78–85.
Stern, C. D. (2005). Neural induction: Old problem, new ﬁndings, yet more questions.
Development, 132, 2007–2021.
Tang, F., Kaneda, M., O’Carroll, D., Hajkova, P., Barton, S. C., Sun, Y. A., et al. (2007). Maternal
microRNAs are essential for mouse zygotic development. Genes & Development, 21, 644–648.
Tawk, M., Araya, C., Lyons, D. A., Reugels, A. M., Girdler, G. C., Bayley, P. R., et al. (2007).
A mirror-symmetric cell division that orchestrates neuroepithelial morphogenesis. Nature, 446,
Valencia-Sanchez, M. A., Liu, J., Hannon, G. J., & Parker, R. (2006). Control of translation and
mRNA degradation by miRNAs and siRNAs. Genes & Development, 20, 515–524.
Viczian, A. S., Solessio, E. C., Lyou, Y., & Zuber, M. E. (2009). Generation of functional eyes
from pluripotent cells. PLoS Biology, 7, e1000174.
Vieira, C., Pombero, A., Garcia-Lopez, R., Gimeno, L., Echevarria, D., & Martinez, S. (2010).
Molecular mechanisms controlling brain development: An overview of neuroepithelial
secondary organizers. The International Journal of Developmental Biology, 54, 7–20.
Principles of Early Vertebrate Forebrain Formation
Wienholds, E., Koudijs, M. J., van Eeden, F. J., Cuppen, E., & Plasterk, R. H. (2003). The
microRNA-producing enzyme Dicer1 is essential for zebraﬁsh development. Nature Genetics,
Wilson, S. W., & Houart, C. (2004). Early steps in the development of the forebrain.
Developmental Cell, 6, 167–181.
Yamaguchi, T. P. (2001). Heads or tails: Wnts and anterior-posterior patterning. Current Biology:
CB, 11, R713–R724.
Zuber, M. E., Gestri, G., Viczian, A. S., Barsacchi, G., & Harris, W. A. (2003). Speciﬁcation of the
vertebrate eye by a network of eye ﬁeld transcription factors. Development, 130, 5155–5167.
Control of Organogenesis by Hox Genes
J. Castelli-Gair Hombría, C. Sánchez-Higueras
and E. Sánchez-Herrero
Abstract Hox genes encode a class of animal transcription factors well known for
the segment transformations they generate when mutated or expressed ectopically.
Hox genes are stably expressed during development in partially overlapping
antero-posterior domains of the body where they impose their morphological
characteristics. This is achieved in two main ways: ﬁrst, Hox proteins are capable of
activating (or repressing) the expression of gene networks responsible for cell
speciﬁcation and organ formation, and second, they compete out the activity of
other Hox proteins, either by transcriptional repression or by posterior prevalence.
Studies in Drosophila indicate that Hox proteins regulate genes required for organ
development, indicating that Hox genes play a role in organogenesis that goes
beyond providing antero-posterior regionalization. In a few cases Hox expression is
transient, and the input is just required for organ speciﬁcation. However, in other
cases the Hox proteins remain active after organ speciﬁcation and their function is
required for fundamental aspects of organogenesis and cell differentiation.
Á Hox Á Gene networks Á Drosophila Á Development
Hox genes encode homeodomain transcription factors that confer speciﬁc morphological characteristics to the regions of the body where they are expressed.
Mutations in Hox genes can cause spectacular homeotic transformations, where one
segment transforms its morphology into that of a neighboring segment. The ﬁrst
J. Castelli-Gair Hombría (&) Á C. Sánchez-Higueras
Centro Andaluz de Biología del Desarrollo (CSIC/JA/Universidad Pablo de Olavide),
Centro de Biología Molecular-Severo Ochoa (CSIC/Universidad Autónoma de Madrid),
© Springer International Publishing Switzerland 2016
J. Castelli-Gair Hombría and P. Bovolenta (eds.), Organogenetic Gene Networks,
J. Castelli-Gair Hombría et al.
Hox mutation described, bx1, was isolated in Drosophila by Calvin Bridges around
1915 and was later studied in depth by Edward B. Lewis, who found it mapped to a
region of the chromosome where other homeotic mutations clustered. Lewis published a comprehensive genetic analysis of this region, named the Bithorax complex
(BX-C), and suggested it contained several genes controlling the morphological
divergence of each thoracic and abdominal segment (Lewis 1978). Later work
revealed that the BX-C is composed of only three genes: Ultrabithorax (Ubx),
abdominal-A (abd-A) and Abdominal-B (Abd-B) (Sánchez-Herrero et al. 1985;
Tiong et al. 1985) and that many of the mutations originally isolated where affecting
cis-regulatory elements regulating the temporal and spatial expression of these three
genes. A second homeotic complex was found, the Antennapedia complex
(ANT-C) that included ﬁve Hox genes specifying the morphology of cephalic and
anterior thoracic segments: labial (lab), proboscipedia (pb), Deformed (Dfd), Sex
combs reduced (Scr) and Antennapedia (Antp) (Kaufman et al. 1980, 1990)
(Fig. 12.1). Lewis proposed that the BX-C originated by gene duplication in an
ancestral segmented millipede-like arthropod with a body composed of identical
repeated units. After duplication, the BX-C genes would have evolved by mutation,
acquiring novel functions that resulted in the stepwise diversiﬁcation of the segment
shape along the anterior-posterior body axis (Lewis 1978). However, molecular
analyses demonstrated that Hox genes are also present in vertebrates and they must
have appeared much earlier in evolution (McGinnis et al. 1984a, b, c; Scott and
Hox genes were originally seen as factors implementing genetic switches
between homologous segments, conferring to each of them a deﬁned genetic
Fig. 12.1 Hox cluster organization in fruit fly, worm and mouse. Hox genes localized in the same
cluster are represented as a box on a continuous line, the color of the box represents gene
homology. The relative position of most Hox genes in the cluster is maintained during evolution
and as a result orthologous genes tend to appear in columns. In Drosophila the single cluster has
split in two. In the Nematode Caenorhabditis elegans, many Hox genes have been lost but the
Abd-B like homolog has experienced an expansion (green boxes). In mice, as in humans, two
cluster duplications have given rise to four Hox clusters (Hox a–Hox d). The Drosophila group 3
genes have evolved losing their Hox function and they are not represented in this ﬁgure. Modiﬁed
from Foronda et al. (2009)
Control of Organogenesis by Hox Genes
address, constant in time and uniform in space. Later research revealed the complex
temporal and spatial control of these genes, their role in elaborating genetic circuits
and their speciﬁc tissue and organ requirements. In this chapter, focusing mostly in
Drosophila, we review the function of Hox genes in organogenesis.
The Origin of Hox Genes
Hox genes can be found in all animals except sponges (Porifera) and comb jellies
(Ctenophora) (Holland 2013). Hox clusters evolved from a smaller primordial
cluster probably containing only four genes, similar to the situation now present in
simple animals like Cnidarians and Acoeles. The number of Hox genes in this
hypothetical cluster expanded by tandem duplication explaining why all existing
Hox genes can be classiﬁed in one of four categories (Fig. 12.1), known as:
Anterior, Group 3, Central or Posterior Hox genes (Garcia-Fernandez 2005a, b).
These duplications gave rise to a cluster formed by seven Hox genes that is likely to
represent the situation at the time when the Cambrian explosion of animal forms
occurred. Afterwards, independent duplications expanded the number of Hox genes
per cluster from 9 to 15 in different animal lineages, while in other lineages there
was a Hox gene loss. Loss is especially evident in Nematode worms, which have
lost up to ﬁve Hox orthologs (Aboobaker and Blaxter 2003). While originally Hox
genes were organized as a single cluster, in some animals the cluster split, as is now
observed in Drosophila melanogaster, where it has subdivided into the ANT-C and
BX-C (Fig. 12.1).
An extreme case of evolution by duplication of whole Hox clusters occurred in
the lineage leading to vertebrates. Cephalochordates have a single Hox cluster,
which is thought to be the primitive Chordate situation, but in vertebrates two
successive whole genome duplication events gave rise to four clusters, named
HoxA, HoxB, HoxC and HoxD in mouse and human. In teleost ﬁsh, additional
whole genome duplication probably led to the existence of eight Hox clusters.
These duplications caused a certain level of redundancy that was followed by Hox
gene losses, leading to a ﬁnal number of seven clusters (Hueber et al. 2010). As a
consequence of these genomic changes, the total number of Hox genes varies from
the 15 Hox genes organized in a single cluster of the Cephalocordates, to the 39
genes in four clusters present in mouse and human and the 46 to 49 Hox genes in
seven clusters found in various ﬁsh (Holland 2013; Garcia-Fernandez 2005a, b;
Aboobaker and Blaxter 2003; Hueber et al. 2010).
Although the large evolutionary distances separating all animal phyla makes it
difﬁcult to establish direct correspondence among Hox genes, their common origin
from an ancient cluster is reflected by the presence of orthologous genes.
Orthologous genes derive from the same gene present in the cluster before the
species diverged (or the whole genome duplications occurred) and thus are more
similar to a gene in another species than to other Hox genes in the cluster where it is
located. Thus, when comparing the human and Drosophila Hox sequences, Hox1