The SP in the CA3 region has a unique U-shaped curve that reaches the dentate hilus. In rats, the HP appears from E18 and expands curvilinearly until E21, at which time the layer begins to medially expand and forms a U-shaped structure by E22 (Altman and Bayer, 1990b). The CA3 pyramidal neurons have a neurogenic gradient such that pyramidal neurons near the CA1 region are generated earlier than those near the dentate gyrus (Bayer, 1980). The neurogenesis gradient from ventral to dorsal is also observed in the CA3 region. The neurogenesis of CA3 pyramidal neurons takes place in the VZ between E16 and E20, with a peak between E17 and E18 in rats, which is earlier than the generation of hippocampal CA1 neurons that peak around E18 and E19 in rats (Bayer, 1980; Altman and Bayer, 1990a,b). The generated neurons move to just above the VZ and transform into a multipolar morphology, similar to CA1 pyramidal neurons (Nakahira and Yuasa, 2005). However, CA3 neurons remain longer in the MAZ than CA1 neurons. In mice, neurons generated in the CA3 region at E14 exhibit multipolar morphology even at E18, while CA1 pyramidal neurons born at the same time have already migrated into the HP (Nakahira and Yuasa, 2005). This phenomenon is also observed in rats using administration of [3H] thymidine (Altman and Bayer, 1990b). Reports have shown that the sojourning period for CA3 pyramidal neurons assuming a multipolar morphology is 1 day longer than for CA1 neurons (Altman and Bayer, 1990b; Nakahira and Yuasa, 2005). Altman and Bayer hypothesized that multipolar cells in the CA3 region might wait for a connection with granule cells of the dentate gyrus, resulting in a longer sojourning time (Altman and Bayer, 1990b). Again, the report by Deguchi et al. showed that neurons generated at the same time in different sub-regions connect preferentially with each other (Deguchi et al., 2011), which may also explain a longer sojourn to wait for a connection with dentate gyrus cells. The CA3 pyramidal neurons accumulate in the MAZ for 4 days then migrate upward. The migration modes of CA3 pyramidal neurons are thought to be similar to CA1 pyramidal neurons, though only a limited number of studies have been reported on the behavior of hippocampal CA3 pyramidal neurons (Nakahira and Yuasa, 2005). Interestingly, Nakahira and Yuasa also found that neurons generated at E16 in the mouse CA3 VZ migrate tangentially toward the subpial area, and that some neurons then detach from the stream and migrate radially, with a unipolar shape, along with radial glial fibers directed to the HP of the CA3 region (Nakahira and Yuasa, 2005). This neuronal behavior is different from CA3 neurons generated at E14. Although the CA3 neurons born at E16 account for only a small portion (Bayer, 1980), multiple migration modes may exist for CA3 neurons depending on their birthdates.

The major difference between the CA1 and the CA3 during hippocampal development is the layer shape. The HP in the CA1 is mildly curved and in parallel with its ventricular surface, whereas the CA3 HP has a U-shape with one end invading the dentate hilus. This end is apart from the VZ. Considering that neurons at this end of the HP are also generated from the VZ in the CA3, the long journey for the migrating cells is thought to be one of the causes of delayed HP formation in the CA3, which occurs 1 day later than HP formation in the CA1 (Altman and Bayer, 1990b).

Although the CA1 and the CA3 are continuous architectures via the CA2, pyramidal cells in each region express specific markers; for example, SCIP, a POU domain transcriptional factor, in CA1 pyramidal neurons (Frantz et al., 1994; Tole et al., 1997), and KA1, a glutamate receptor subunit, in CA3 neurons (Wisden and Seeburg, 1993; Tole et al., 1997). Interestingly, these markers are already expressed in each cell group at E15.5 in mice, when the cells are still localized in the IZ (Tole et al., 1997). Explant culture experiments performed by Tole et al. revealed that this marker is expressed in a cell-autonomous manner (Tole and Grove, 2001). Therefore, the destination of cells comprising the HP in the CA3 is thought to already be determined at, or soon after, the multipolar cell stage. For future CA3 pyramidal cells, especially those in the HP end of the dentate hilus, the climbing migration mode may be adequate for migration and detours to reach their final positions. Eventually, CA3 pyramidal neurons might migrate more horizontally through the HP than CA1 pyramidal neurons.

# Migration of Cells Comprising the Dentate Gyrus

Since Altman found postnatal neurogenesis in the subgranular zone (SGZ) of the dentate gyrus (Altman, 1963), it is wellestablished that the dentate gyrus is one of the two regions where adult neurogenesis occurs. Compared with studies on adult neurogenesis, the development of the dentate gyrus and migratory dynamics of granule cells have been less extensively studied. The generation and migration of dentate cells during development are complex and quite different from pyramidal neurons in the Ammon's horn (**Figure 2**). Dentate cells are generated in the primary dentate neuroepithelium located around the dentate notch, which is ventral to the Ammonic VZ and dorsal to the fimbria at E16 and E17 of rats (Altman and Bayer, 1990c) and at E13.5 and E14 in mice (Li et al., 2009; Seki et al., 2014). By E15.5 in mice (E18 in rats), some cells that are generated in the primary dentate VZ (Altman and Bayer, 1990c; Seki et al., 2014) migrate out to the subpial region through the suprafimbrial region, which is

FIGURE 2 | Schema of migration of dentate cells during hippocampal development. Newborn granule cells from the dentate notch migrate to the secondary dentate matrix (SDM) (indicated by a brown arrow). The cells then migrate to the subpial surface to form the outer part of the dentate granule cell layer (light blue arrow 1), followed by the dentate hilus (light blue arrow 2), which is called the tertiary dentate matrix (TDM) at this stage, to later form the inner part of the layer. Cells in the TDM exhibit proliferative activities into adulthood, although the proliferative region becomes restricted to the subgranular zone. In contrast, pyramidal neurons in the hippocampal CA1 and neocortex are generated in the Ammonic ventricular zone and the neocortical ventricular zone, respectively, and migrate in a radial direction (indicated by magenta and black arrows, respectively).

designated as "dentate migration." During this migration, these cells still exhibit proliferative activity and form the secondary dentate matrix (SDM in **Figure 2**) on the migratory route (Altman and Bayer, 1990c; Pleasure et al., 2000). On the migrating stream, the cells extend many processes to various directions (Nakahira and Yuasa, 2005), and the migration route of granule cells and progenitors are separated into two routes. One is the "first dentate migration" in which early-born cells migrate to the crest of the dentate gyrus through the subpial route (route 1 in **Figure 2**) (Altman and Bayer, 1990c). The dentate cells in the subpial region also exhibit highly proliferative activity (Li et al., 2009). The morphologies of cells in the subpial region are diverse; for example, bipolar morphology and multipolar cell morphology (Nakahira and Yuasa, 2005). This diversity is probably owing to the variety of cellular maturation. The cells reaching the subpial region radially migrate toward the supra-granular blade of the dentate gyrus apposed to the radial glial fibers (Nakahira and Yuasa, 2005). At this stage, these cells form a unipolar or bipolar shape with one or two branches (Nakahira and Yuasa, 2005). The dentate granule cells in the subpial region first form the outer shell of the supra-granular blade of the dentate gyrus, and then gradually shift to form the outer shell of the infra-granular blade of the dentate gyrus. Around E17.5 and 18.5 in mice, a prototype of the dentate gyrus can be observed (Seki et al., 2014). In late embryonic and early postnatal days, the "second dentate migration" appears in which the cells migrate toward and reach the future dentate hilus (route 2 in **Figure 2**). Because these cells still exhibit proliferative activity even in the hilus, the zone where these cells exist is called the tertiary dentate matrix (TDM in **Figure 2**). Dentate neurons generated from the TDM form the inner part of the dentate gyrus. As a result, dentate granule cells in the dentate granule cell layer adopt a birthdate-dependent outside-in pattern, in which earlier-born neurons locate to the outer part of this layer and later-born neurons position to the inner region (Rakic and Nowakowski, 1981; Altman and Bayer, 1990c). Although neurogenesis in the TDM continues into adulthood, this zone becomes gradually restricted to the boundary between the dentate granule cell layer and the hilus, called the SGZ (Altman and Bayer, 1990c). Bayer suggests that granule cells in the dentate gyrus are generated from E15 to adulthood in rats, and about 80–85% of total granule cells are generated after birth (Bayer, 1980).

Seki et al. traced the granule cells using a glial fibrillary acidic protein (GFAP)-GFP transgenic mouse line (Seki et al., 2014). GFAP was not thought to be a marker for "embryonic" progenitors of dentate granule cells, although GFAP is a well-known marker for "adult" progenitors. Seki et al. found that migrating progenitors of dentate granule cells, unlike those of neocortical pyramidal neurons, express GFAP from the beginning of dentate development, and these cells could be traced using a GFAP-GFP transgenic mouse line. Immunohistochemical analyses using this transgenic mouse line showed maturation of granule cells during migration. For example, neurogenin-positive proneural cells are mainly localized in the VZ, whereas Tbr2-positive early neural progenitors are principally located in the migratory stream and the developing dentate gyrus. NeuroD-positive immature neurons are mostly located in the migratory stream, the developing dentate gyrus, and the hilus, whereas prox1-positive granule cells are positioned in the developing dentate gyrus and the hilus. Sox2-positive progenitor cells are distributed all over the dentate gyrus at E18 in mice. Accordingly, while these granule cells gradually mature during migration, cells in each region are heterogeneous as to their degree of maturation. How this heterogeneous group could migrate along the same route is not yet known.

# Molecular Mechanism of Neuronal Migration in the Hippocampus

A number of mutant mouse lines or the introduction of shRNAexpression vectors for various genes into neurons in utero have been used to show abnormal layer formation and mis-positioning of neurons in the Ammon's horn and the dentate gyrus during hippocampal development. Some of the examples are summarized below.

# Reelin (and Related Molecules, ApoER2, VLDLR, and Dab1)

Reelin is a giant glycoprotein secreted from Cajal–Retzius cells in the MZ during development (D'Arcangelo et al., 1995; Hirotsune et al., 1995; Ogawa et al., 1995).  For example, in the neocortex, the reelin-deficient autosomal recessive mouse, reeler, displays disrupted layer formation, including overall approximate inversion of the birthdate-dependent layering (Caviness, 1973). Anatomical analyses, such as[3H] thymidine injection and in situ hybridization, disclosed that hippocampal layer formation is also inverted in the reeler mouse (Caviness, 1973; Stanfield and Cowan, 1979a,b; Stanfield et al., 1979; Niu et al., 2004; Boyle et al., 2011). Injection of CR-50, a function-blocking antibody against Reelin protein, into the ventricle of mouse embryos also resulted in a similar layer pattern to that of reeler mice in the hippocampal Ammon's horn (Nakajima et al., 1997). Additionally, reeler mice exhibit a divided SP in the CA1, a meandered SP in the CA3, a less densely packed dentate gyrus, and a reduced number of granule cells (Caviness, 1973; Stanfield and Cowan, 1979a,b; Boyle et al., 2011).

Reelin binds to Apolipoprotein E receptor 2 (ApoER2) and very low-density lipoprotein receptor (VLDLR) (D'Arcangelo et al., 1999; Hiesberger et al., 1999; Trommsdorff et al., 1999), subsequently induces phosphorylation of Disabled-1 (Dab1) by Fyn or Src kinases, and then transduces the signal to several downstream pathways to regulate neuronal migration and cellular positioning (Honda et al., 2011; Sekine et al., 2014). Double KO mice of apoer2 and vldlr show similar phenotypes to those of reeler mice. Additionally, KO mice of apoer2 show slightly more severe SP splitting than the vldlr KO mice, while the phenotypes of these single KO mice are milder than those of double KO mice (Trommsdorff et al., 1999; Drakew et al., 2002; Weiss et al., 2003). The dab1 KO mice also exhibit the same hippocampal abnormality as the reeler mice (Howell et al., 1997; Weiss et al., 2003).

Analysis of the migratory stream of dentate cells using nestin-GFP transgenic mice mated with reeler mice suggests that Reelin is not necessary for the migration of dentate precursors (Nestin-GFP positive cells) from the dentate notch to the subpial zone, but is indispensable for later migration of these cells from the subpial zone to the granule cell layer (Li et al., 2009). Prox1-positive granule cells in these mice are arranged abnormally in the dentate area, suggesting the involvement of Reelin in the final radial migration of dentate granule cells. Forster et al. reported that Reelin and Dab1 affect radial glial cell differentiation and branching in the hippocampus and the loss of these genes causes abnormal radial fiber formation, resulting in failed neuronal migration in the hippocampus (Förster et al., 2002). Zhao et al. rescued reeler malformation in the orientation of radial glial fibers and dentate cell migration in vitro during dentate gyrus development when they adjacently co-cultured reeler dentate gyrus with rat wild-type dentate gyrus (Zhao et al., 2004, 2006). These results collectively imply that Reelin controls neuronal migration in the hippocampus in cell-autonomous and non-cell-autonomous manners.

# Cyclin-dependent Kinase 5 (Cdk5)/p35

Cdk5 is a serine/threonine kinase and is activated by binding with its regulatory co-factor p35 or p39. Cdk5 activity is rich in brains during development, and p35 or p39 expressions are restricted to the brain. Cdk5 regulates various aspects of neuronal functions such as cell migration, cytoskeletal remodeling, and adult neurogenesis via phosphorylation of various types of molecules (Su and Tsai, 2011). The cdk5 KO mice are lethal by birth and display disrupted laminar formation in the neocortex and hippocampus (Ohshima et al., 1996). The p35 KO mice also exhibit mis-positioning of neuronal cells in the Ammon's horn (partly distinct SP is observed in the CA3, but not in the CA1) and the dentate gyrus, though the phenotypes are moderate compared with cdk5 KO mice (Wenzel et al., 2001; Ohshima et al., 2005). Cdk5 regulates multipolar-to-bipolar transition of migratory neurons in the neocortex via RapGEF2 phosphorylation (Ohshima et al., 2007; Ye et al., 2014). Because hippocampal pyramidal neurons also perform this transition, a similar mechanism may also play a role during Ammon's horn development.

# Doublecortin (Dcx)

DCX is a microtubule-associated protein involved in neuronal migration and a causative gene for X-linked lissencephaly (des Portes et al., 1998; Gleeson et al., 1998). Patients with lissencephaly with DCX mutations exhibit defects in neocortical and hippocampal lamination (Barkovich et al., 1991). Hemizygous male dcx mutant mice are lethal by early postnatal days and display disrupted hippocampal formation, that is, abnormal neuronal positioning/migration in the Ammon's horn and partial dividing of the SP in the CA3, while neocortical laminar formation and the dentate gyrus are quite normal (Corbo et al., 2002). The dcx heterozygous female mice display milder malformation in the hippocampus and deficits in learning and memory (Corbo et al., 2002). KO mice of dclk1 or dclk2, doublecortin-like kinase 1 or 2, respectively, are anatomically normal in the hippocampus (Deuel et al., 2006; Tanaka et al., 2006; Kerjan et al., 2009), but the double KO mice of dcx and dclk1 show severe abnormalities in hippocampal laminar formation in the Ammon's horn and the dentate gyrus (Deuel et al., 2006; Tanaka et al., 2006). In contrast, double KO mice of dcx and dclk2 exhibit disrupted laminar formation in CA1 and CA3 region and a less-packed dentate granule layer (Kerjan et al., 2009). Considering the majority of mutant mice mentioned in this review also show abnormalities in the neocortex, the hippocampus and neocortex are likely to share molecular pathways during development. Mutant mice that exhibit malformations specifically in the hippocampal region, such as dcx KO mice, may become a key tool to better understand the molecular mechanisms underlying the unique process of hippocampal development, such as the climbing mode of migration. Analyses of dcx KO mice and double KO mice of dcx and dclk1 or dclk2 may also provide insight into differences between hippocampal CA1 and CA3 regions.

# Pafah1b1 (formerly Lis1)

PAFAH1B1 is another causative gene for lissencephaly (Reiner et al., 1993; Hattori et al., 1994; Lo Nigro et al., 1997). Pafah1b1 regulates microtubule-based transport by binding with Dynein motor proteins and Ndel1 (formerly Nudel) (Sasaki et al., 2000). Heterozygous Pafah1b1 KO mice exhibit malformations of the hippocampal cytoarchitecture, which results from delayed neuronal migration and abnormal cellular positioning (Hirotsune et al., 1998; Fleck et al., 2000). Consequently, hippocampal layers in the Ammon's horn become discontinuous and multiple in this mouse, whereas granule cells in the dentate gyrus are less concentrated and loosely packed (Fleck et al., 2000). Another lissencephaly-associated protein, tubulin alpha 1A (Tuba1a), is also involved in hippocampal layer formation (Keays et al., 2007). The tuba1a S140G mutant mice induced by injection of N-ethyl-N-nitrosourea (ENU) exhibit deficits in neuronal migration, resulting in a double layer of the hippocampal CA1 and CA3 regions, as well as abnormal laminar formation in the neocortex (Keays et al., 2007).

# Cxcl12 (SDF-1)/Cxcr4

SDF-1 is another secreted protein that regulates cellular migration (Bleul et al., 1996; Ma et al., 1998; Klein et al., 2001). In the dentate gyrus, SDF-1 is expressed in the meninges and Cajal–Retzius cells, whereas its receptor Cxcr4 is expressed in migratory granule cells in the second dentate matrix and the migratory stream (Bagri et al., 2002). Disruption of the normal SDF-1 gradient by the ectopic SDF-1 expression to the hippocampal field using electroporation into slice culture causes a deficit in granule cell migration, suggesting SDF-1 is a chemoattractant factor for dentate migration (Bagri et al., 2002). The cxcr4 KO mice exhibit a disrupted dentate gyrus, caused by migration defects of granule cells along the subpial stream and subsequent radial migration (Bagri et al., 2002; Lu et al., 2002; Li et al., 2009).

# Nuclear Factor Ib (Nfib)

Nfib is a member of nuclear factors I (Nfia, b, c, and d) and functions as a transcriptional factor. The nfib KO mice display abnormal hippocampal formation, including the CA3, dentate gyrus, and fimbria, which may be due to aberrant maturation of radial glial fibers in the Ammon's horn (Steele-Perkins et al., 2005; Barry et al., 2008). Although the deficient mouse exhibits normal cell proliferation, the dentate cells accumulate in the subpial region during dentate migration. Barry et al. suggest that this abnormal positioning of dentate cells is attributed to failed radial migration (Barry et al., 2008).

### Disrupted-in-Schizophrenia-1 (Disc1)

DISC1 is a risk gene for major psychiatric disorders, including schizophrenia. We have previously reported that Disc1 knockdown by in utero electroporation causes abnormal migration of hippocampal CA1 neurons; knockdown cells fail to enter the pyramidal layer (Tomita et al., 2011). Disc1 is also related to migration and positioning/integration of dentate granule cells during development and adulthood (Duan et al., 2007; Kim et al., 2009; Meyer and Morris, 2009). KO mice of girdin, a Disc1 interacting molecule, show dispersion of granule cells in the dentate gyrus, mis-positioning of pyramidal neurons in the CA1 and the CA3, and split layer in the CA2 (Enomoto et al., 2009). Knockdown of girdin in dentate granule cells results in overmigration, as observed for the Disc1 knockdown cells. Because other genes implicated in neuropsychiatric diseases, such as CNT-NAP2 (Peñagarikano et al., 2011) and FMRP (La Fata et al., 2014), are reported to be involved in neuronal migration during neocortical development, it will be interesting to determine whether these genes also affect hippocampal neuronal migration.

Hippocampal neuronal migration is controlled by extracellular factors, such as Reelin, SDF-1, and radial glial fibers, as well as intracellular molecules as mentioned above. Both the reeler mouse and the nfib KO mouse exhibit abnormal radial glial fibers and disrupted dentate gyrus formation. However, both KO mice exhibit normal dentate cell migration and cells accumulate in the subpial region; the final migration toward the dentate layer is conducted along radial fibers, whereas dentate migration to the subpial region may be independent of these molecules.

Pafah1b1, Dcx, and Tuba1A regulate microtubule dynamics. Dab1 is reported to bind to Lis1 downstream of Reelin signaling (Assadi et al., 2003). Dcx is phosphorylated by Cdk5 at Ser297, resulting in reduced microtubule polymerization and binding affinity to microtubules (Tanaka et al., 2004). Cdk5 also regulates the Pafah1b1-Ndel1-Dynein complex via Ndel1 phosphorylation (Niethammer et al., 2000; Sasaki et al., 2000). Furthermore, Cdk5 and Reelin signaling synergistically contribute to neuronal positioning (Ohshima et al., 2001, 2007; Beffert et al., 2004). Therefore, microtubule dynamics is critical for migration of the hippocampal cells, similar to neocortical neurons.

A number of mutant mice exhibiting abnormal hippocampal formation also display splitting of the SP in the Ammon's horn, although the extent of splitting is not uniform. This may suggest the existence of multiple migration modes for hippocampal pyramidal neurons. In the rodent hippocampus, deep and superficial sublayers are visibly distinguished by cellular density and morphology in the ventral two-thirds of CA1 (Slomianka et al., 2011). Slomianka et al. also reviewed the distinction of histological, molecular, and connective features between deep and superficial sublayers in the hippocampal Ammon's horn (Slomianka et al., 2011). For example, superficial pyramidal cells express Satb2 during development, Nov and Nr3c2 in the hippocampal CA1 in adulthood, and Kcnq5 in the CA3 (Thompson et al., 2008; Dong et al., 2009). In contrast, deep pyramidal neurons express Sox5 during development, Ndst4 and Astn2 in the CA1 in adulthood, and St18 in the CA3 (Thompson et al., 2008; Dong et al., 2009). Furthermore, Mizuseki et al. showed physiological differences between deep and superficial sublayers in the hippocampal CA1 region of rats, such as theta phase preference during REM sleep and gamma phase preference during behavioral task (Mizuseki et al., 2011). In the hippocampal CA1, the climbing mode of migration is observed for late-born neurons (Kitazawa et al., 2014). In contrast, Morest reported that hippocampal cells extended their leading process through the HP and kept it until they reach the pial surface during the early developmental stages of opossum (Morest, 1970). Rodent hippocampal cells may also use this somal-translocation-like mode, especially during early stages of development. If this is the case, early-born neurons and late-born neurons might use different modes of migration and ultimately settle in their respective positions in the SP. The distinct populations between superficial and deep layers may be reflected by specific gene expressions and functions.

# Conclusion

This review discusses the migration of pyramidal neurons in the Ammon's horn and granule cells in the dentate gyrus during hippocampal development. The structure of the hippocampus is dynamically expanded and becomes complicated during development. Because the climbing mode of migration is a flexible migration mode, it may be necessary for hippocampal neurons to accommodate to this hippocampal formation. The migration of dentate cells is well-organized, while cellular maturation is diverse along the migratory stream. Integration of molecular biological studies with histological studies has led to novel discoveries focused on cellular and molecular mechanisms of hippocampal development. Further studies on behaviors of hippocampal neurons are expected in the future to fully understand hippocampal development and functions.

# Acknowledgments

We thank members of the Nakajima Laboratory for valuable discussions. This work was supported by grants from the Strategic Research Program for Brain Sciences ("Understanding of molecular and environmental bases for brain health") (10036057), and the Grants-in-Aid for Scientific Research of the Ministry of Education, Culture, Sports, Science, and Technology of Japan (22111004, 25640039, 25116522, 26430075, 15H01586, 15H02355, 15H01293, 15K09723, 15K18363, 15K06745), SEN-SHIN Medical Research Foundation, Terumo Life Science Foundation, Life Science Foundation of Japan, Keio Gijuku Academic Development Funds, Keio Gijuku Fukuzawa Memorial Fund for the Advancement of Education and Research, and Program for the Advancement of Keio Gijuku Next Generation Research Projects.

# References

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Hayashi, Kubo, Kitazawa and Nakajima. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# **Genotype-phenotype correlation in neuronal migration disorders and cortical dysplasias**

#### *Mitsuhiro Kato\*†*

*Department of Pediatrics, Yamagata University Faculty of Medicine, Yamagata, Japan*

Neuronal migration disorders are human (or animal) diseases that result from a disruption in the normal movement of neurons from their original birth site to their final destination during early development. As a consequence, the neurons remain somewhere along their migratory route, their location depending on the pathological mechanism and its severity. The neurons form characteristic abnormalities, which are morphologically classified into several types, such as lissencephaly, heterotopia, and cobblestone dysplasia. Polymicrogyria is classified as a group of malformations that appear secondary to post-migration development; however, recent findings of the underlying molecular mechanisms reveal overlapping processes in the neuronal migration and post-migration development stages. Mutations of many genes are involved in neuronal migration disorders, such as *LIS1* and *DCX* in classical lissencephaly spectrum, *TUBA1A* in microlissencephaly with agenesis of the corpus callosum, and *RELN* and *VLDLR* in lissencephaly with cerebellar hypoplasia. *ARX* is of particular interest from basic and clinical perspectives because it is critically involved in tangential migration of GABAergic interneurons in the forebrain and its mutations cause a variety of phenotypes ranging from hydranencephaly or lissencephaly to early-onset epileptic encephalopathies, including Ohtahara syndrome and infantile spasms or intellectual disability with no brain malformations. The recent advances in gene and genome analysis technologies will enable the genetic basis of neuronal migration disorders to be unraveled, which, in turn, will facilitate genotype-phenotype correlations to be determined.

**Keywords: lissencephaly, heterotopia, polymicrogyria, tubulinopathy, interneuronopathy,** *LIS1, DCX, ARX*

# **Introduction**

The characteristic six-layered neocortex in the human brain is formed by two types of neuron, projection neurons and interneurons, which migrate from their birth places, such as the ventricular zone and ganglionic eminence, respectively. Neuronal migration disorders are human (or animal) diseases that result from the disruption of normal movement of neurons from their original birth site to their final destination during early development. As a consequence, the neurons remain somewhere along their migratory route, their location depending on the pathological mechanism and its severity. Many genes have been found to be responsible for neuronal migration disorders, such as *LIS1* and *DCX* in classical lissencephaly spectrum, *TUBA1A* in lissencephaly with cerebellar hypoplasia, *ARX* in X-linked lissencephaly with abnormal genitalia (XLAG), *FLNA* and *ARGEF2* in periventricular heterotopia, *FCMD* and glycosylation-related

#### *Edited by:*

*Chiaki Ohtaka-Maruyama, Tokyo Metropolitan Institute of Medical Science, Japan*

#### *Reviewed by:*

*Brian Key, Asdasdasd, Austria Masaharu Hayashi, Tokyo Metropolitan Institute of Medical Science, Japan*

#### *\*Correspondence:*

*Mitsuhiro Kato, Department of Pediatrics, Yamagata University Faculty of Medicine, 2-2-2 Iida-nishi, Yamagata 990-9585, Japan<EMAIL_ADDRESS>
#### *† Present Address:*

*Mitsuhiro Kato, Department of Pediatrics, Showa University School of Medicine, Tokyo, Japan*

#### *Specialty section:*

*This article was submitted to Neurogenesis, a section of the journal Frontiers in Neuroscience*

*Received: 01 December 2014 Accepted: 06 May 2015 Published: 21 May 2015*

#### *Citation:*

*Kato M (2015) Genotype-phenotype correlation in neuronal migration disorders and cortical dysplasias. Front. Neurosci. 9:181. doi: 10.3389/fnins.2015.00181* genes, such as *POMT1*, *POMT2*, *POMGNT1*, *POMGNT2*, *FKRP*, *LARGE*, *TMEM5*, *POMK*, *ISPD*, *GMPPB*, *B3GNT1*, and *B3GALNT2* in cobblestone dysplasias, *GPR56*, *SRPX2*, and some tubulin-related genes, e.g., *TUBA8*, *TUBB2B*, and *TUBB3*, in polymicrogyria (Kato and Dobyns, 2003; Vuillaumier-Barrot et al., 2012; Buysse et al., 2013; Stevens et al., 2013; Fry et al., 2014). Recently, we found that mutations in *COL4A1*, which encodes type IV collagen alpha 1 subunit, cause schizencephaly accompanied by polymicrogyria in the adjacent cortex of the transmantle cleft as well as focal cortical dysplasia (Yoneda et al., 2013). Historically, brain malformations including neuronal migration disorders have been classified based on a postmortem examination. The advancement and spread of neuroimaging techniques, particularly magnetic resonance imaging (MRI), make it easier to find out many types of brain malformations, but make it more complicated to classify them. Moreover, the unveiling of responsible genes for brain malformations has changed the classification scheme and causes most neuroscientists and even physicians trouble to follow it. Here, I review the clinical manifestation of neuronal migration disorders, focusing mainly on lissencephaly, in terms of genotype-phenotype correlations.

# **Lissencephaly Spectrum: Classical Lissencephaly to Subcortical Band Heterotopia**

Lissencephaly is classified as a spectrum of disorders caused by widespread abnormal transmantle migration, ranging from classical lissencephaly (agyria or pachygyria) to subcortical band heterotopia or double-cortex syndrome (Barkovich et al., 2012). Classical lissencephaly is characterized by a smooth (*lissos* in Greek) brain surface with a decreased number of sulci and wide gyri. Mutations in *LIS1*, located on chromosome 17p13.3, or *DCX* on Xq23 are the main cause for classical lissencephaly (**Table 1**) (Kato and Dobyns, 2003). Mutations in *DCX* are causative for classical lissencephaly in male individuals and subcortical band heterotopia in female individuals. A combination of a severity grading scale [the most severe form, Grade 1 (total agyria) to the mildest form, Grade 6 (subcortical band heterotopia) via the intermediate forms comprised of a combination of agyria, pachygyria, and subcortical band heterotopia] and an anterior or posterior gradient scale is useful to predict the causative gene for lissencephaly spectrum (Kato and Dobyns, 2003). For instance, mutations of *LIS1*, *ARX*, or *TUBA1A* result in a posterior more severe than anterior gradient, while mutations of *DCX* or *RELN* lead to an anterior more severe than posterior gradient. LIS1 participates in cytoplasmic dynein-mediated nucleokinesis, somal translocation, and cell motility (Smith et al., 2000) as well as mitosis or neurogenesis and chromosomal segregation (Faulkner et al., 2000). DCX is a microtubule-associated protein and is involved in microtubule polymerization and stabilization (Gleeson et al., 1999). Missense mutations in *DCX* responsible for lissencephaly spectrum are mainly located in two tandem repeats (N-terminal or C-terminal doublecortin domains), which bind to microtubules or free tubulin and other components (Friocourt et al., 2005), respectively.

 Agyria is generally characterized by the disappearance of deep sulci in more than one lobe and the thickness of the cortex is 10–20 mm (**Figure 1**). The gyri in pachygyria are wider than in the normal cortex and the thickness of the cortex is 4–9 mm (**Figure 2**). Brain MRI of subcortical band heterotopia shows bilateral continuous symmetric bands of gray matter underlying an almost normal cortical mantle with relatively shallow sulci (**Figure 3**).  Subcortical band heterotopia in male patients is caused by somatic mosaic *DCX* mutations or *LIS1* mutations (Gleeson et al., 2000; Kato et al., 2001; D'agostino et al., 2002; Poolos et al., 2002). Coexistence of agyria and pachygyria or pachygyria and subcortical band heterotopia can be seen in the same patient, suggesting common mechanisms for these phenotypes. Microscopically, agyria and pachygyria present a four-layered cortex with an outer molecular layer, superficial layer, cell sparse layer, and deep cellular layer. In the marginal zone between pachygyria and subcortical band heterotopia, the outer molecular layer corresponds to layer I of the normal sixlayered cortex, the superficial layer corresponds to layers II–VI, the cell sparse layer corresponds to subcortical white matter, and the deep cellular layer corresponds to band heterotopia with a mass of unlayered ectopic neurons (**Figure 4**). The primary pathology of lissencephaly due to the *DCX* mutations shows only minor differences compared with that caused by *LIS1* mutations, for example, inferior olivary ectopia is present in *LIS1* mutation brains but is absent in *DCX* mutation brain (Berg et al., 1998); however, Viot et al. report a different cortical architecture for *DCX* lissencephaly (Viot et al., 2004).

The severity of the clinical manifestations of lissencephaly spectrum is correlated with the degree of brain malformation. Patients with agyria show severe muscle hypotonia from infancy (known as floppy infant) and achieve neither head control nor are they able to say meaningful words. A specific form of epileptic seizure, epileptic spasms, occurs in 80% of patients with agyria or pachygyria, although electroencephalography (EEG) may not present with typical hypsarrhythmia, which is characteristically seen in infantile spasms or West syndrome (Guerrini, 2005). However, the main clinical features of subcortical band heterotopia are intellectual disability and epileptic seizures, both of which are milder than those of agyria or pachygyria. Intellectual disability ranges from normal to severe retardation and correlates with the thickness of the band and the degree of pachygyria (Barkovich et al., 1994; Bahi-Buisson et al., 2013). Genetic counseling is particularly important for parents that have a boy with classical lissencephaly or a girl with subcortical band heterotopia because the mother may be a heterozygous carrier of the *DCX* mutation.

Miller-Dieker syndrome is a contiguous gene syndrome caused by a microdeletion in 17p13.3, a region that contains *LIS1* and *YWHAE* (which encodes 14-3-3 protein epsilon). Phenotypes of Miller-Dieker syndrome are more severe than

*pontocerebellar hypolasia; PMG, polymicrogyria.*

**157**

**FIGURE 1 | Complete agyria in a** *DCX* **mutation patient (Grade 1 on the severity scale).** T2-weighted axial MRI image. Wide shallow sylvian fissures create a figure-of-eight appearance. The thickness of the cortex is over 10 mm. A high-intensity (white) line (arrow heads) beneath the cerebral surface is consistent with a cell sparse layer of the four-layered cortex.

that of classical lissencephaly because of an isolated *LIS1* mutation. They are characterized by complete agyria and facial abnormalities including prominent forehead, bitemporal hollowing, short nose with upturned nares, prominent upper lip with downturned vermilion border and small jaw, and sometimes other congenital defects involving the heart, kidneys, intestine, or fingers (Kato and Dobyns, 2003). Neurological findings of Miller–Dieker syndrome are similar to those of patients with agyria, such as severe developmental delay with weak muscle tone and profound intellectual disability, intractable seizures,

**FIGURE 3 | Subcortical band heterotopia or double cortex syndrome in a** *DCX* **mutation patient (Grade 5 on the severity scale).** T2-weighted axial MRI image. Subcortical heterotopic gray matter in the posterior region fuses into the pachygyric cortex in the anterior region (arrowheads).

dysphagia, and poor prognosis with recurrent infection of the respiratory system.

# **Tubulin-Related Disorders, Tubulinopathies**

Microtubules provide the main structural framework for the shafts of axons and dendrites, and with actin serve as tracks for intracellular trafficking and to provide the driving force underlying neurite extension and intracellular movement of organelles during mitosis (Flynn et al., 2013).  All the above are classified as tubulinopathies (Cushion et al., 2013; Bahi-Buisson et al., 2014). Microtubules are assembled from soluble tubulin heterodimers consisting of alpha- and betatubulin. Multiple isoforms of both tubulins are encoded by different genes. Mutations of *TUBA1A*, which encodes alpha tubulin, cause lissencephaly spectrum, particularly diffuse agyria or perisylvian pachygyria, with microcephaly, agenesis of the corpus callosum, and cerebellar hypoplasia (**Figure 5**) (Bahi-Buisson et al., 2008). *TUBA1A* mutations account for only 1% of isolated classical lissencephaly; however, they account for approximately 30% of patients with lissencephaly associated with cerebellar hypoplasia (Kumar et al., 2010). Dysgenesis of the anterior limb of the internal capsule and disorganization of the hippocampus are other neuroimaging features for *TUBA1A* mutation (Poirier et al., 2007). Mutations of *TUBA1A* cause polymicrogyria as well. Interestingly, mutations of *TUBB2B* cause polymicrogyria with or without congenital fibrosis of the external ocular muscles as well as bilateral perisylvian pachygyria(Cederquist et al., 2012; Romaniello et al., 2014). Polymicrogyria is classified as a group of malformations that appear secondary to post-migration development; however, recent findings of the underlying molecular mechanisms reveal overlapping process in neuronal migration and post-migration development stages.

Mutations of *TUBA8* cause polymicrogyria with optic nerve hypoplasia and display autosomal recessive inheritance (Abdollahi et al., 2009). Mutations of *TUBB2A*, which encodes beta-tubulin, cause infantile-onset epilepsy with simplified gyral patterning (Cederquist et al., 2012; Cushion et al., 2014; Romaniello et al., 2014). Mutations of *TUBB3* cause two distinct forms. One is congenital fibrosis of the external ocular muscles or oculomotor nerve hypoplasia and later-onset peripheral axon degeneration with dysgenesis of the corpus callosum, anterior commissure, and internal capsule, but with no cortical dysplasia suggesting migrational defects (Tischfield et al., 2010). Another is cortical dysgenesis including polymicrogyria, pontocerebellar hypoplasia, and abnormal basal ganglia, but with no ocular motility defects (Poirier et al., 2010). The main mechanisms underlying the phenotypes caused by*TUBB3* mutations are impaired axon guidance owing to disrupted microtubule dynamics and kinesin interaction (Tischfield et al., 2010). Tubulinopathies caused by the mutations of the genes encoding alpha- or beta-tubulin demonstrate more extensive phenotypes compared to other gene mutations, such as *LIS1*, *DCX*, or *RELN*. Mutations of *TUBA1A*, which encodes alphatubulin 1A, is the most frequently found in patients with brain malformations, while more genes encoding beta-tubulin, such as *TUBB2A*, *TUBB2B*, *TUBB3*, *TUBB4A*, and *TUBB*, are identified in a wide spectrum of disorders besides brain malformations. Pathological mechanisms and discrepancy between alpha- and beta-tubulinopathies should be elucidated.

# **ARX-Related Disorders, Interneuronopathies**

The embryonic cerebral cortex at the stage of neuronal migration contains neuronal cells with two modes of migration; radial migration from the ventricular zone toward the pia and tangential migration from ganglionic eminence along a tangential trajectory into the developing cortex. Radially

The boundary of the caudate nucleus and lentiform nucleus is obscure. Complete agenesis of the corpus callosum and pontocerebellar hypoplasia are also seen.

migrating neurons in the cortex are mainly excitatory projection neurons expressing glutamate as a neurotransmitter. Tangentially migrating neurons are inhibitory interneurons expressing the neurotransmitter GABA. XLAG is caused by mutation of *ARX*, which is expressed in the embryonic ganglionic eminence, neocortex, and hippocampus and plays important roles in neuronal proliferation, interneuronal migration, and differentiation in the embryonic forebrain, as well as a secondary role in differentiation of the testes (Kitamura et al., 2002). Patients with XLAG present occipital-predominant classical lissencephaly, particularly anterior pachygyria and posterior agyria, or a simplified gyral pattern, agenesis of the corpus callosum, and abnormal basal ganglia (Kato et al., 2004). In the most severe form of XLAG, patients show hydranencephaly with a large occipital cavity. Female carriers of *ARX* mutations causing XLAG have a risk of agenesis of the corpus callosum with no cortical defects. Abnormalities of external genitalia range from hypoplastic penis or undescended testes to complete female appearance, while the karyotype is 46,XY. Neuropathological studies show a complete loss or a decreased number of cortical interneurons in human XLAG and in *Arx*-null mice (Bonneau et al., 2002; Kitamura et al., 2002) and a three-layered cortex in human XLAG (Forman et al., 2005). Patients with XLAG show intractable seizures soon after birth, suggesting a great disparity between excitatory projection neurons and inhibitory interneurons. *ARX* mutations in patients with XLAG are null mutations or non-conservative missense mutations at critical amino acids in the homeodomain, while other missense mutations or expansion mutations in the polyalanine tract result in X-linked intellectual disability with or without dystonia, West syndrome, Ohtahara syndrome, or early infantile epileptic encephalopathy with suppression burst on EEG but with no brain malformation (Bienvenu et al., 2002; Stromme et al., 2002; Guerrini et al., 2007; Kato et al., 2007, 2010). Interestingly, longer polyalanine expansion is correlated with more severe and earlier onset phenotypes. A wide spectrum of ARX-related disorders forms a group of interneuronopathies based on the role of ARX during neurogenesis, as seen in patients and in the *Arx*-null mouse model (Kato and Dobyns, 2005; Marsh et al., 2009).

# **Classical Lissencephalies Associated with Other Forms of Brain Malformation**

Classical lissencephaly caused by *LIS1* or *DCX* mutations usually exist in isolated forms and only show cortical dysplasia on brain MRI. Rare variant forms of lissencephaly are associated with congenital microcephaly, cerebellar hypoplasia, or agenesis of the corpus callosum. Each form demonstrates characteristic radiological findings and some of the causative genes have been identified.

A lissencephaly group with cerebellar hypoplasia can be classified into several types according to brain imaging, additional clinical features, and causative genes (Ross et al., 2001). Among them, frontal predominant mild lissencephaly (diffuse pachygyria) with severe hippocampal and cerebellar hypoplasia or Reelin-type lissencephaly is caused by mutation of either *RELN* or *VLDLR* and shows autosomal recessive inheritance (Hong et al., 2000; Boycott et al., 2005). Dysequilibrium syndrome is an allelic disorder of the *VLDLR* locus (Moheb et al., 2008). Reelintype lissencephaly has an inverted or no clear pattern of cortical lamination attributable to abnormal migration of the neurons in an outside-in birth order (Cooper, 2008; Dekimoto et al., 2010).

Lissencephaly can be associated with congenital microcephaly, though the head circumference of lissencephaly caused by the *LIS1* or *DCX* mutations is usually within the normal range. Lissencephaly with a head circumference of less than −3 SD at birth is classified as microlissencephaly (Barkovich et al., 2005) or microcephaly with lissencephaly (Barkovich et al., 2012). Although many genes identified to be responsible for primary microcephaly, such as *MCPH1*, *ASPM*, *CENPJ*, *CDK5RAP2*, and *PNKP*, are involved with the cell-cycle phase of mitosis affecting neurogenesis (Barbelanne and Tsang, 2014), the causative genes for microlissencephaly remain unknown in many cases. Mutations of *WDR62*, which encodes a protein localized to centrosomes throughout mitosis and nucleoli during interphase, cause microcephaly with pachygyria or polymicrogyria (Bilguvar et al., 2010). Mutations of *NDE1*, which encodes a protein that binds dynein and functions in centrosome duplication, as well as the *TUBA1A* mutations mentioned above, cause microcephaly with a simplified gyral pattern, agenesis of the corpus callosum, and cerebellar hypoplasia (Alkuraya et al., 2011; Bakircioglu et al., 2011).

# **Conclusion**

Neuronal migration disorders are classified based on causative genes as well as on brain MRI and neuropathological findings. There are strong relationships between clinical manifestations and mutation of a particular gene, in accordance with the expression and functions of that gene. Recent advances in gene and genome analysis technology will enable the genetic basis of neuronal migration disorders to be readily determined, facilitating the elucidation of genotype-phenotype correlations.

# **Acknowledgments**

The author is grateful to the patients and their families for their kind cooperation. This study was partially funded by the Ministry of Health, Labour and Welfare of Japan (26310301), and an Intramural Research Grant (24-7, 27-5) for Neurological and Psychiatric Disorders from the National Center of Neurology & Psychiatry. This study was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

# **References**

**Conflict of Interest Statement:** The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Copyright © 2015 Kato. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*

# Neuronal migration abnormalities and its possible implications for schizophrenia

#### Kazue Muraki and Kenji Tanigaki\*

*Shiga Medical Center, Research Institute, Moriyama, Shiga, Japan*

#### Edited by:

*Chiaki Ohtaka-Maruyama, Tokyo Metropolitan Institute of Medical Science, Japan*

#### Reviewed by:

*Luca Bonfanti, University of Turin, Italy Emanuel DiCicco-Bloom, University of Medicine and Dentistry of New Jersey/Robert Wood Johnson Medical School, USA*

#### \*Correspondence:

*Kenji Tanigaki, Shiga Medical Center, Research Institute, Moriyama 5-4-30, Shiga 524-8524, Japan<EMAIL_ADDRESS>
#### Specialty section:

*This article was submitted to Neurogenesis, a section of the journal Frontiers in Neuroscience*

> Received: *30 November 2014* Paper pending published: *27 December 2014* Accepted: *20 February 2015* Published: *10 March 2015*

#### Citation:

*Muraki K and Tanigaki K (2015) Neuronal migration abnormalities and its possible implications for schizophrenia. Front. Neurosci. 9:74. doi: 10.3389/fnins.2015.00074* Schizophrenia is a complex mental disorder that displays behavioral deficits such as decreased sensory gating, reduced social interaction and working memory deficits. The neurodevelopmental model is one of the widely accepted hypotheses of the etiology of schizophrenia. Subtle developmental abnormalities of the brain which stated long before the onset of clinical symptoms are thought to lead to the emergence of illness. Schizophrenia has strong genetic components but its underlying molecular pathogenesis is still poorly understood. Genetic linkage and association studies have identified several genes involved in neuronal migrations as candidate susceptibility genes for schizophrenia, although their effect size is small. Recent progress in copy number variation studies also has identified much higher risk loci such as 22q11. Based on these genetic findings, we are now able to utilize genetically-defined animal models. Here we summarize the results of neurodevelopmental and behavioral analysis of genetically-defined animal models. Furthermore, animal model experiments have demonstrated that embryonic and perinatal neurodevelopmental insults in neurogenesis and neuronal migrations cause neuronal functional and behavioral deficits in affected adult animals, which are similar to those of schizophrenic patients. However, these findings do not establish causative relationship. Genetically-defined animal models are a critical approach to explore the relationship between neuronal migration abnormalities and behavioral abnormalities relevant to schizophrenia.

Keywords: schizophrenia, GABAergic interneuron, mouse models, Neuregulin1, DISC1, 22q11 deletion syndrome, Dgcr8, Cxcr4

# Introduction

Schizophrenia is a chronic psychiatric disorder with a strong genetic component. Twin studies indicated that the heritability for schizophrenia is estimated to be 0.81 (Sullivan et al., 2012). Most of genetic linkage studies failed to identify highly-shared risk alleles due to the complexity of genetic architecture of schizophrenia except for DISC1 (Millar et al., 2000). Many combinations of different gene variants cause genetic risk of schizophrenia. Genome-wide association studies have identified many schizophrenia susceptibility candidate genes. Most of such common variants confer only slight increase in risk for schizophrenia (odds ratio < 1.2) (Ripke et al., 2013), and often failed to be replicated. Some of them, Neuregulin1, ErbB4, and Reelin are involved in the regulation of neuronal migration. On the other hand, rare and de novo chromosomal microdeletion or microduplication [copy number variations (CNVs)] have been implicated in schizophrenia (Levinson et al., 2011; Rees et al., 2014). Such CNVs are from thousands to millions nucleotides and contain many genes and their odds ratios are high (2∼20) compared with common variants. 22q11.2 deletion syndrome (22q11DS) is the most frequent known genetic cause of schizophrenia (Pulver et al., 1994). However, it remains to be elucidated how combinations of these genetic variants play pathogenic roles of schizophrenia.

Schizophrenia is believed to result from embryonic developmental abnormalities not from neuronal degenerations (Weinberger, 1987; O'Connell et al., 1997). Cytoarchitectural abnormalities were reported in the entorhinal cortex (Jakob and Beckmann, 1986; Arnold et al., 1991, 1997; Falkai et al., 2000) and the subcortical white matters in schizophrenia (Akbarian et al., 1993a,b, 1996). Decreased neuronal density in the superficial white matter and increased density in the deep white matter suggest neuronal migration defects in schizophrenia. However, other studies failed to replicate these findings (Akil and Lewis, 1997; Krimer et al., 1997; Bernstein et al., 1998; Beasley et al., 2002, 2009), suggesting these abnormalities might be too subtle to be detected without special methods. Prior human brain imaging studies also have indicated reduced cerebral volume, ventricular enlargement, and reduced hippocampal volume in schizophrenia (Shenton et al., 2001).

The development of the human cerebral cortex is similar to that in the mouse (Rakic, 2009; Hansen et al., 2013; Ma et al., 2013), which enabled investigation of the functions of schizophrenia susceptibility candidate genes in neuronal developments. The mammalian cerebral cortex consists mainly of excitatory glutametergic and inhibitory GABAergic neurons. Glutamatergic neurons are generated from neural progenitors in the dorsal forebrain (Glover et al., 2009; Rakic, 2009), whereas most of inhibitory neurons are thought to derive from the ventral pallium: medial, lateral and caudate ganglionic eminence (MGE, LGE, and CGE) (Hansen et al., 2013; Ma et al., 2013). Two types of migration are observed in the cortex. One is radial migration of glutamatergic neurons from the underlying ventricular zone along the radial glial fiber, while the other is tangential migration from the ventral forebrain of GABAergic interneurons (Corbin et al., 2001; Marin and Rubenstein, 2003). However, the differences between the human and mouse cortices are also reported. In the mouse, about 70% of cortical interneurons generate from the MGE and ∼30% are from the CGE (Miyoshi et al., 2010). In contrast, more than half of interneurons derive from the CGE in the human (Hansen et al., 2013). The human cortex showed a much higher diversity in the interneuron types compared with that of the rodents (Feldman and Peters, 1978). In spite of the limitations caused by these differences, animal models are still valuable to elucidate the roles of neuronal migration deficits in the pathogenesis of schizophrenia. Many genetically-modified animal models with construct validity and cell-specific gene modification technique are available. The great advantage of rodent model to study schizophrenia is that we can establish a causal relationship between genetic abnormalities, neuronal developments, and behavioral abnormalities. Here we review a group of studies using rodent models which give insights into the pathogenesis of schizophrenia.

# Developmental Neuronal Disruption Model of Schizophrenia

Perinatal insult of neuronal development can cause anatomical and behavioral deficits similar to human schizophrenic patients. One of the examples is a gestational day 17 (GD17) methylazoxymethanol acetate (MAM) administration rat model (Grace and Moore, 1998; Flagstad et al., 2004; Gourevitch et al., 2004; Paredes et al., 2006). MAM is a mitotic toxin and MAM administration specifically disrupts proliferating region. GD17 MAM treatment results in specific subtle reductions in the volume of prefrontal cortex (PFC) and hippocampus (HP), heterotopias in the HP resulting from neuronal migration deficits (Le Pen et al., 2006; Moore et al., 2006), which are characteristics of schizophrenia (Kovelman and Scheibel, 1984; Shenton et al., 2001; Heckers, 2004; Honea et al., 2005). GD17 MAM-treated rats also display decreased density of parvalbumin (PV)-positive interneurons in medial PFC and HP (Lodge et al., 2009). Interestingly, postmortem studies of human schizophrenia have shown decreased expression of PV and the 67 KDa isoform of glutamic acid decarboxylase (GAD67), which is an enzyme responsible for GABA synthesis, in PFC of schizophrenia subjects (Akbarian et al., 1995; Volk et al., 2000; Hashimoto et al., 2003; Fung et al., 2010). However, no difference is observed in the density of PV-positive interneurons in schizophrenia (Hashimoto et al., 2003). PVpositive interneurons are known to be indispensable for synchronized firing of excitatory pyramidal neurons in gamma frequencies (30–80 Hz), which plays essential roles for cognitive functions (Howard et al., 2003). Altered gamma oscillation activity and cognitive deficits have been reported in schizophrenia (Cho et al., 2006; Minzenberg et al., 2010). Consequently, PV-positive interneuron deficits are thought to be the cause of impairments of gamma oscillation and cognition in individuals with schizophrenia (Lewis et al., 2012). MAM-treated rats also show behavioral deficits in prepulse inhibition (PPI), which reflects an inability to filter out irrelevant sensory information, and working memory task (Flagstad et al., 2005; Le Pen et al., 2006; Moore et al., 2006), which are typical symptoms of schizophrenia in humans (Braff and Geyer, 1990; Liddle and Morris, 1991; Goldman-Rakic, 1994; Swerdlow et al., 1994; Nuechterlein et al., 2004). Furthermore, electrophysiological studies have shown that enhanced activity of ventral HP leads to dopaminergic neuronal activation in MAMtreated rats (Lodge and Grace, 2007). Again, these altered hippocampal activities are also observed in human schizophrenic patients (Medoff et al., 2001; Schobel et al., 2009). These abnormalities can be normalized by administration of α5GABA A receptor positive allosteric modulator, SH-053-2′F-R-CH3 (Gill et al., 2011), suggesting the involvement of GABA in embryonic MAM treatment-induced deficits. The MAM model provides a direct evidence that subtle embryonic disruptions of neuronal development result in behavioral alterations disorders, although the etiology is absolutely different from that of schizophrenia in humans.

# Neuregulin-ERBB Signaling

Neuregulins are a large family of epidermal growth factor (EGF)-like proteins and play divergent roles both in neuronal development and in the neuronal activity homeostasis in the mature central nervous system. Several genetic linkage studies have shown Neuregulin1 (NRG1) as a strong candidate gene for schizophrenia (Badner and Gershon, 2002; Stefansson et al., 2002; Lewis et al., 2003). Some GWASs also support the hypothesis (Li et al., 2006; Munafo et al., 2006; Shi et al., 2009; Agim et al., 2013), although it has not been confirmed by a recent mega-analysis in which international consortia combined the resources to maximize the sample size and identified more than 100 candidate genes for schizophrenia with high levels of statistical significance (Schizophrenia Working Group of the Psychiztric Genomics Consortium, 2014). Most of schizophreniaassociated single nucleotide polymorphisms (SNPs) in NRG1 are localized in the 5′ and 3′ region of the gene. Some of them are associated with the expression level of NRG1 (Law et al., 2006; Weickert et al., 2012). A receptor of NRG1, ERBB4 is also associated with schizophrenia (Benzel et al., 2007; Law et al., 2007; Shi et al., 2009; Agim et al., 2013). It has been reported that a rare chromosome micro-deletion in a schizophrenic patient disrupts this gene, resulting in a truncated protein similar to dominant-negative ERBB4 (Walsh et al., 2008).

Nrg1 generates six types and at least 30 isoforms owing to multiple promoters and alternative splicing (Mei and Nave, 2014). Most pro-Nrg1 isoforms are transmembrane proteins and N-terminal domains containing EGF-like domain are released out of the cell after undergoing proteolytic processing except for TypeIII Nrg1 (cysteine-rich-domain containing Nrg1 (CRD-Nrg1)). This released mature Nrg1 activates ErbB receptor tyrosine kinase such as ErbB2/ErbB3 heterodimer and ErbB4 homodimers. Nrg1 regulates migration of excitatory glutamatergic neurons and γ-aminobutyric acid (GABA)-producing interneurons in the embryonic cortex. Nrg1 promotes the maintenance of radial glial cells in the cortex and induces elongation of radial fiber, which are essential for the radial migration of cortical excitatory neurons and cerebellar granule cells (Anton et al., 1997; Rio et al., 1997). NRG1 is also critical for interneuronal tangential migration (Flames et al., 2004; Li et al., 2012). ErbB4 is expressed in interneuronal progenitors migrating from the MGE to the cortex (Yau et al., 2003; Flames et al., 2004). Type III Nrg1 is expressed in lateral ganglionic eminence, and Type I and II Nrg1 (immunoglobulin (Ig)-domain containing Nrg1 (Ig-Nrg1)) are expressed in the embryonic cortex (Flames et al., 2004). Diffusible Type I and II Nrg1 in the cortex are thought to attract ErbB4 expressing interneurons along a permissive corridor of Type III Nrg1 (Flames et al., 2004), although this model is challenged. In another model, Nrg1, and Nrg3 have been proposed to be repellants for migrating interneurons (Li et al., 2012). Loss of ErbB4 cuases embryonic lethality due to failed development of myocardial trabeculae, which made it difficult to characterize the functions of ErbB4 signaling in interneuronal migration (Gassmann et al., 1995; Kramer et al., 1996). However, heart-rescued ErbB4 knockout mice with cardiac-specific ErbB4 transgene displayed decreased number of GABAergic interneurons in the postnatal cortex (Flames et al., 2004; Fisahn et al., 2009; Li et al., 2012), which clearly showed the essential roles of Nrg1/ErbB4 signaling in interneuronal migration.

Cell-specific gene modification techniques are now starting to elucidate a link between Nrg1/ErbB4 signaling and pathophysiology of schizophrenia. The gain and loss of function of Nrg1/ErbB4 signaling were examined because postmortem studies of schizophrenia reported both increased and decreased NRG1/ERBB4 signaling in schizophrenic patients (Silberberg et al., 2006; Law et al., 2007; Weickert et al., 2012; Joshi et al., 2014). Transgenic mice overexpressing Type I Nrg1 showed deficits in PPI and contextual fear conditioning, and hyperlocomotion (Deakin et al., 2009, 2012; Yin et al., 2013; Luo et al., 2014). If the overexpression of Nrg1 was switched off in adult mice, its effects were reversible (Yin et al., 2013; Luo et al., 2014). The influence of Nrg1 overexpression on neuronal development remains to be elucidated.  Nrg1 heterozygous mice and heart-rescued ErbB4 knockout (KO) mice showed decreased number of cortical PV interneurons (Fisahn et al., 2009; Neddens and Buonanno, 2010; Shamir et al., 2012; Pei et al., 2014) (**Table 1**). However, PV interneuron-specific deletion of ErbB4 did not affect the number of cortical interneurons, which might be due to the slow turnover of ErbB4 (Fazzari et al., 2010; Shamir et al., 2012) (**Table 1**). A comparative behavioral analysis of ErbB4 KO and PV interneuron-specific ErbB4 KO mice demonstrated that PV interneuron-specific deletion is sufficient for hyperactivity and deficits in PPI. The only difference is that ErbB4 KO mice but not PV interneuron-specific ErbB4 KO mice exhibit reduced anxiety-like behaviors and deficits in cued and contextual fear conditioning (Shamir et al., 2012), which might be caused by developmental disorders in ErbB4-deficient interneurons.

# Disrupted-in Schizophrenia 1

The disrupted-in schizophrenia 1 (DISC1) gene was discovered at the breakpoint of inherited balanced chromosomal translocation in a Scottish family suffering from major depression, schizophrenia, and bipolar disorder (St Clair et al., 1990; Millar et al., 2000). Following linkage analysis and association studies demonstrated that DISC1 is significantly associated with schizophrenia, bipolar disorder and major depression (Ekelund et al., 2001, 2004; Macgregor et al., 2004; Hamshere et al., 2005; Hashimoto et al., 2006; Liu et al., 2006; Thomson et al., 2014), although it has not been confirmed by a recent mega-analysis of GWASs (Schizophrenia Working Group of the Psychiztric Genomics Consortium, 2014).

DISC1 is a scaffolding protein interacting with multiple proteins: nuclear distribution gene E homolog-like 1 (NDEL1), lissencephaly-1 (LIS1), phosphodiesterase 4B (PDE4B), glycogen

*OF, open field, PPI:prepulse inhibition; FC, fear conditioning; TM, transmembrane region; PV, parvalbumin; HC, hippocampus; ND, not determined.*

synthase kinase 3β (GSK3β) and DIX domain-containing 1 (DIXDC1) (Ozeki et al., 2003; Millar et al., 2005; Taya et al., 2007; Mao et al., 2009). DISC1/DIXDC1 functions as a switch between neuronal proliferation and migration. DISC1/DIXDC1 binds to GSK3β and inhibits its activity leading to proliferation of neural progenitors through the inhibition of β-catenin degradation (Mao et al., 2009). CDK5 phosphorylation of DISC1 at S710 and DIXDC1 facilitates neuronal migration by dissociating DISC1 from GSK3β and promoting its binding with NDEL1 (Singh et al., 2010; Ishizuka et al., 2011). DISC1 variants associating with human brain structures and psychiatric phenotypes have been reported to impair this switching mechanism (Singh et al., 2011). Furthermore, knockdown of DISC1 also impairs interneuronal tangential migrations (Steinecke et al., 2012, 2014).

Two hypotheses are proposed on the pathophysiology of the disruption of the DISC1 gene: that the Scottish mutation decreases DISC1 expression and leads to haploinsufficiency; or that the Scottish mutation results in production of carboxyterminal-truncated DISC1 (amino acids 1-598). This C-terminal truncated DISC1 functions as a dominant negative protein, and impairs microtubule dynamics by blocking interaction between DISC1 and dynein complex (Kamiya et al., 2005). Dynein complex contains LIS1 and NDEL1, and regulates coupling of the nucleus and centrosome, which is indispensable for radial migration of cortical excitatory neurons (Sasaki et al., 2000). Knockdown of DISC1 inhibits cortical neuronal cell migration (Kamiya et al., 2005; Kubo et al., 2010).

Acute knockdown of DISC1 using RNAi leads to drastic neuronal migration deficits. However, DISC1 KO mice (Disc112-3/12-3) astonishingly showed almost normal cytoarchitectures of the cerebral cortex and the HP (Kuroda et al., 2011), whereas the number of PV-positive interneurons reduced in female Disc112-3/12-3 mice (Nakai et al., 2014). The phenotypes in the proliferation of neuronal progenitors have not been examined in DISC1 KO mice, which will provide important insight into the pathogenesis of DISC1 deficiency. Furthermore, Disc112-3/12-3 mice did not show schizophrenia-like phenotype but exhibited lower anxiety and higher impulsivity (Kuroda et al., 2011) (**Table 2**). Only female Disc112-3/12-3 mice exhibited enhanced responsiveness to methamphetamine and deficits in PPI (Kuroda et al., 2011). These milder phenotypes of Disc112-3/12-3 mice might be explained by a compensation mechanism after chronic loss of DISC1. N-nitroso-N-ethylurea (ENU) mutagenesis was utilized to generate missense mutations of Disc1. L100P mutant (Disc1L100P/L100P) mice showed reduced brain volume, reduced number of cortical neurons, altered distribution of cortical neurons, interneuonal migration deficits, and schizophrenia-like behavioral abnormalities, although the behavioral phenotypes were not confirmed by another group due to the difference in the genetic background (Clapcote et al., 2007; Lee et al., 2011, 2013; Shoji et al., 2012). 129S6/SvEv 25-bp deletion variant results in the production of a truncated isoform of DISC1 (amino acids 1-542) (Koike et al., 2006). C57BL/6J mice carrying the Disc1 gene from the 129S6/SvEv strain (Disc1125 bp/125 bpmice) exhibited enlarged ventricle and working memory deficits (Koike et al., 2006; Juan et al., 2014). Neuron-specific overexpression of the truncated DISC1 also resulted in drastic phenotypes: enlarged lateral ventricle, reduced number of PV-positive interneurons and schizophrenia-like behavioral abnormalities (Hikida et al., 2007; Pletnikov et al., 2008; Shen et al., 2008; Ayhan et al., 2011) (**Table 2**). Furthermore, a technique of inducible transgene expression enabled a specific expression of truncated DISC1 during only prenatal period, only postnatal period or both periods (Ayhan et al., 2011). Prenatal expression only led to decreased brain volume and decreased number of PV-positive interneurons. Enlarged lateral ventricle seems to be affected by postnatal expression of truncated DISC1. In contrast, enhanced responsiveness to psychostimulant required prenatal and postnatal continuous expression (Ayhan et al., 2011), which suggests that both neurodevelopmental abnormality and neuronal functional impairment caused by truncated DISC1 might be essential for pathogenesis of schizophrenia.

# 22q11 Deletion Syndrome

22q11.2 deletion syndrome (22q11DS) is the most frequent known genetic cause of schizophrenia (Pulver et al., 1994). 22q11DS accounts for about 1% ofschizophrenia cases (Karayiorgou et al., 1995; Manolio et al., 2009). Prior brain imaging studies of human 22q11 DS have indicated reduced cerebral volume, ventricular enlargement and reduced hippocampal volume (Eliez et al., 2000, 2001; Chow et al., 2002; Simon et al., 2005). All of these brain anomalies have also been reported in schizophrenia (Shenton et al., 2001). All of the genes except for one gene in human 22q11.2 locus exist on mouse chromosome 16 (Puech et al., 1997). This has facilitated the generation of mouse models of 22q11 DS, which carry a hemizygous deletion of 22q11-related region of mouse chromosome16 (Lindsay et al., 1999; Paylor and Lindsay, 2006; Stark et al., 2008). These animal models show schizophrenia-related behavioral abnormalities such as working memory deficits, sensory information-processing deficits, and enhanced responsiveness to psychostimulants (Paylor et al., 2001; Stark et al., 2008; Earls et al., 2011; Kimoto et al., 2012), which are recognized as major deficits of schizophrenia (Elvevag and Goldberg, 2000; Green et al., 2000; Swerdlow et al., 2001). Animal models of 22q11DS showed reduced density of layer II– IV projection neurons in a medial PFC (Meechan et al., 2009), reduced volume of a perinatal HP dentate gyrus (Toritsuka et al., 2013), delayed migration of hippocampal dentate neuronal progenitors and cortical interneurons, and altered distribution of PV-positive interneurons (Meechan et al., 2009, 2012; Toritsuka et al., 2013), although it remains to be elucidated these deficits in neurogenesis lead to excitatory/inhibitory imbalance or not. Perinatal hippocampal DG and interneuronal migration abnormalities are caused by Cxcl12/Cxcr4 signaling deficits (Toritsuka et al., 2013). Cxcl12/Cxcr4 signaling might play pivotal roles in the pathogenesis of schizophrenia. Previous studies also suggest a possible involvement of Cxcl12/Cxcr4 signaling in the neurodevelopmental disorders of GD17 MAM-treated animal model of schizophrenia (Paredes et al., 2006). The expression of CXCL12 is decreased in olfactory neurons from sporadic cases with schizophrenia compared with normal controls (Toritsuka et al., 2013).

Among of genes deleted in 22q11DS, Dgcr8 is a promising candidate gene for schizophrenia-related phenotypes. Dgcr8 forms the microprocessor complex of microRNA (miRNA) with Drosha, which is essential for miRNA production. Overexpression of Dgcr8 rescued interneuronal migration deficits of 22q11DS model mice, and the migration of hippocampal DG and interneuronal progenitors were also affected inDgcr8+/<sup>−</sup> mice (Toritsuka et al., 2013). These observations demonstrated the important roles of Dgcr8 in the pathogenesis of 22q11DS. miRNA–mediated regulation network fine tunes the balance of signaling and confers robustness to the system (Herranz and Cohen, 2010). miRNA-mediated regulation can buffer increases or reductions in gene dosage (Staton et al., 2011). Haplodeletion of Dgcr8 causes 20–70% reduction of a specific subsets of mature miRNAs both in Dgcr8 heterozygous and 22q11DS model mice (Stark et al., 2008). Dgcr8 heterozygousity might uncover the effects of 22q11 microdeletion through the disruption of miRNA-mediated buffering effects. In mice, heterozygous deletion of Dgcr8 alone showed working memory

deficits, sensory information-processing deficits and some of neurodevelopmental abnormalities such as reduced cortical neuronal densities (Stark et al., 2008; Fenelon et al., 2011). The behavioral abnormalities and neurodevelopmental disorders of Dgcr8+/<sup>−</sup> mice are similar but some of them are milder than those of 22q11DS model mice,(Stark et al., 2008; Meechan et al., 2009; Fenelon et al., 2011), which might suggest that additional haplodeletion of other genes in 22q11-related regions might be required for the complete reconstitution of phenotypes of 22q11DS model mice. It remains to be elucidated whether behavioral abnormalities of 22q11DS model mice are directly caused by neuronal migration deficit and Cxcr4 signaling defects.

# References

# Concluding Remarks

Elucidating the relationship between neurodevelopmental abnormalities and the pathogenesis of schizophrenia would be exceptionally difficult. In order to dissect the complex causal relations, more sophisticated genetic manipulation would be required. Combination of various techniques such as conditional knockout, inducible transgene expression and virus-mediated gene delivery will enable cell type-specific and developmental stagespecific knockout or rescue experiments. In the future comprehensive profile of neurodevelopmental deficits-behavioral abnormalities will provide significant insights into mental disease pathogenesis of all these neurodevelopmental genes.

subset of prefrontal cortical gamma-aminobutyric acid neurons in subjects with schizophrenia. Arch. Gen. Psychiatry 57, 237–245. doi: 10.1001/archpsyc.57.3.237

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2015 Muraki and Tanigaki. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Genetic manipulation of reptilian embryos: toward an understanding of cortical development and evolution

# *Tadashi Nomura1,2\*, Wataru Yamashita3, Hitoshi Gotoh1 and Katsuhiko Ono1*

*<sup>1</sup> Developmental Neurobiology, Kyoto Prefectural University of Medicine, Kyoto, Japan*

*<sup>2</sup> Japan Science and Technology Agency, PRESTO, Kawaguchi, Japan*

*<sup>3</sup> Department of Biophysics, Graduate School of Science, Kyoto University, Kyoto, Japan*

#### *Edited by:*

*Chiaki Ohtaka-Maruyama, Tokyo Metropolitan Institute of Medical Science, Japan*

#### *Reviewed by:*

*Loreta Medina, Universidad de Lleida, Spain Alino Martinez-Marcos, Universidad de Castilla, Spain Tomomi Shimogori, RIKEN Brain Science Institute, Japan*

#### *\*Correspondence:*

*Tadashi Nomura, Developmental Neurobiology, Kyoto Prefectural University of Medicine, INAMORI Memorial Building, Shimogamo-hangi cho 1-5, Sakyo-ku, Kyoto 606-0823, Japan e-mail<EMAIL_ADDRESS>
The mammalian neocortex is a remarkable structure that is characterized by tangential surface expansion and six-layered lamination. However, how the mammalian neocortex emerged during evolution remains elusive. Because all modern reptiles have a homolog of the neocortex at the dorsal pallium, developmental analyses of the reptilian cortex are valuable to explore the origin of the neocortex. However, reptilian cortical development and the underlying molecular mechanisms remain unclear, mainly due to technical difficulties with sample collection and embryonic manipulation. Here, we introduce a method of embryonic manipulations for the Madagascar ground gecko and Chinese softshell turtle. We established *in ovo* electroporation and an *ex ovo* culture system to address neural stem cell dynamics, neuronal differentiation and migration. Applications of these techniques illuminate the developmental mechanisms underlying reptilian corticogenesis, which provides significant insight into the evolutionary steps of different types of cortex and the origin of the mammalian neocortex.

**Keywords: amniotes, reptiles, cortex,** *in ovo* **electroporation,** *ex vivo* **culture, evolution**

# **INTRODUCTION**

The mammalian cerebral cortex is a remarkable brain structure that is responsible for intricate social behaviors and intelligence. The cerebral cortex is characterized by tangential expansion of its surface area, which is particularly enhanced in the primate and human neocortex, and a six-layered laminar structure composed of multiple types of excitatory and inhibitory neurons (Nieuwenhuys, 1994; Kriegstein et al., 2006; Defelipe, 2011; Lui et al., 2011). The basic frameworks of these unique characteristics are accomplished by the dramatic increase in the number of neural stem/progenitor cells and massive irruption of distinct types of neurons, followed by the coordinated migration of differentiated neurons during embryogenesis. Recent advances of developmental neurobiology have illuminated the molecular mechanisms that govern these complicated cellular events during corticogenesis (Campbell, 2005; Flames and Marin, 2005; Dehay and Kennedy, 2007; Molyneaux et al., 2007; Kumamoto and Hanashima, 2014).

On the contrary, the origin and evolutionary process of the mammalian cortex remain elusive. Phylogenic and paleontological evidence indicated that the forerunners of the mammalian lineage diverged from the common ancestors of amniotes at approximately 300 million years ago (Carroll, 1988; Ruta et al., 2003, 2013). Other lineages of amniotes have also diverged into several unique animal groups that include the descent of extant reptiles (Ruta et al., 2003). In recent years, numerous fossil records have been identified from Paleozoic and Mesozoic sediments, which provided significant information on the process of amniote diversification. Three-dimensional tomographic analyses of fossil endocasts suggested that the size of the mammalian cerebral cortex has increased rapidly in accordance with the dependence of olfactory and somatosensory information (Quiroga, 1979; Rowe et al., 2011); however, histological architectures of the ancestral cerebral cortex remains unknown, preventing us from tracing how the cerebral cortex has specifically evolved in the mammalian lineage.

Ontologically, the cerebral cortex is derived from the dorsal pallium (DP), which develops in the dorsal part of the telencephalon in all vertebrate species (Northcutt, 1981; Puelles et al., 2000; Cheung et al., 2007; Aboitiz, 2011). Despite of developmental homology to the cerebral cortex, the DP in non-mammalian amniotes forms in distinct manners: a three-layer lamination is constructed in the reptilian DP, whereas nuclear slabs are formed in the avian DP (Medina and Reiner, 2000; Heyers et al., 2003; Jarvis et al., 2005; Striedter, 2005). Phylogenetically, aves are included in reptiles (Nomura et al., 2013b; Xu et al., 2014), but here we will use the term reptiles to specifically mean "non-avian reptiles" that include lizards, geckoes, turtles and crocodiles. Because reptiles occupy a unique evolutionary position within amniotes, developmental analyses of the reptilian cortex illuminate commonalities and divergence of developmental programs, thus providing significant insights into the origin of the mammalian cerebral cortex. Previous studies identified unique features of reptilian corticogenesis, such as an outsidein pattern of neuronal migration (Goffinet et al., 1986, 1999; Tissir et al., 2003; Aboitiz and Zamorano, 2013), a difference of layer-specific cell types produced in the reptilian dorsal pallium (Reiner, 1991, 1993), a difference regarding the existence of intermediate progenitors (Charvet et al., 2009; Medina and Abellan, 2009), and lower rates of neurogenesis compared to mouse and other mammalian species (Nomura et al., 2013a). However, modern experimental techniques have not been applied to the analyses of reptilian corticogenesis, largely because of several technical difficulties in collection and manipulation of embryos. First, most reptilian species exhibit seasonal reproduction; thus, a large number of embryos at the desired stages are not constantly available. For example, common lizards/geckoes such as *Lacerta trilineata, Anolis carolinensis*, or *Eublepharis macularius* are frequently used as a model animal in comparative developmental biology (Goffinet et al., 1986; McLean and Vickaryous, 2011; Eckalbar et al., 2012; Sanger et al., 2012). The females of these species produce a limited number of eggs after bleeding. Second, unlike chicken, most reptilian species lay soft-shell eggs, which hampers *in ovo* manipulation of embryos. Although a few pioneering works have reported *in ovo* gene delivery or *ex ovo* culture with snake, lizard and turtle embryos (Nagashima et al., 2007; Matsubara et al., 2014; Tschopp et al., 2014), detailed protocols on embryonic manipulation for reptiles have not been provided.

Here, we describe a method of embryonic manipulation techniques for two reptilian species: the Madagascar ground gecko (*Paroedura pictus*) and the Chinese softshell turtle (*Pelodiscus sinensis*). Surgical techniques on developing reptilian embryos enable us to utilize various experimental approaches. We established the introduction of exogenous genes into the reptilian cortex by *in ovo* electroporation. Furthermore, we developed an *ex ovo* culture system for gecko and turtle embryos, which remarkably increased accessibility to the embryos and improved the efficiency of gene introduction. Successful manipulation techniques of non-mammalian embryos are valuable for studies of the evolutionary developmental biology of the cerebral cortex.

### **ANIMALS AND DETAILED PROTOCOLS OF NEW TECHNIQUE MADAGASCAR GROUND GECKO**

The Madagascar ground gecko (*P. pictus*) is a ground crawling gecko that commonly lives on Madagascar Island. Gecko, lizard, snakes and Tuatara (Sphenodon) are included in the group of lepidosaurs (**Figure 1A**). The adult size of *P. pictus* is approximately 15–20 cm, and the gecko is easy to handle and breed in captivity (**Figure 1B**). After mating of a pair of male and female geckoes, a female produces 1 or 2 clutches every 10–20 days and continues to lay eggs for several months (Nomura et al., 2013a,b). Embryonic staging of the gecko has been established by Noro et al., who determined that the gecko embryogenesis proceeds much slower than chicken embryogenesis (**Figure 1D**) (Noro et al., 2009; Wise et al., 2009). The gecko does not exhibit temperature-dependent sex determination. The embryo hatches approximately 60 days after oviposition and begins to catch small insects within a few days after the first molting. To feed the geckoes, various sizes of crickets were purchased from a local breeder (Tsukiyono farm, Gunma, Japan) and dusted with mineral supplements (calcium and vitamin D) to prevent rickets. To collect embryos, 4 pairs of wild-type geckoes (total 8 animals) were first obtained from a local store (Kansai Reptile Pro, Osaka, Japan) and maintained in our laboratory (28◦C, 12 h of light and dark cycles, 50–60% humidity). More than 100 eggs were obtained from 4 females bred for 6 months.

hindlimb bud becomes larger than forelimb bud, 4: autopodium develops discrete paddle shape, 5: zeugopodium and stylopodium become distinct, 6: digits develop, 9: phalanges develop, 10: claws develop, 11: scale formation and pigmentation, and 12: hatching. Detailed staging criteria are described in Wise et al. (2009).

#### **CHINESE SOFTSHELL TURTLE**

Turtle embryos have been used for anatomical and developmental studies since the nineteenth century (Tokita and Kuratani, 2001). The Chinese softshell turtle (*P. sinensis*) is a freshwaterliving turtle that is widely distributed in eastern and southeastern Asia (**Figure 1C**). The adult size of the turtle reaches over 30 cm in carapace length, and sexual maturity takes approximately 5–6 years. Because the turtle exhibits seasonal reproduction, we could obtained fertilized eggs from a local breeder (Daiwa-Yoshoku, Saga, Japan) in the summer from the beginning of June to the end of August. Sex determination is not dependent on temperature. The developmental stages of the turtle have been established by a previous report (Tokita and Kuratani, 2001). Embryogenesis takes approximately 60 days, and newborn turtles begin foraging after consuming the remaining abdominal yolk. All experimental procedures for reptilian captivity and embryonic manipulation were approved by the experimental animal committee of Kyoto Prefectural University of Medicine (M23-272), and were performed in accordance with the relevant guidelines of the committee.

#### **MANIPULATION AND ELECTROPORATION OF REPTILIAN EMBRYOS**

Embryonic manipulation and electroporation are based on the procedures for gene transduction into developing avian embryos with slight modification (**Figure 2**) (Nomura et al., 2008; Nakamura, 2009). However, because the reptilian eggs are much smaller than chicken eggs, *in ovo* manipulation of reptilian embryos requires specific experimental techniques and training of surgical skills under the dissecting microscope (**Figure 2A**).

**FIGURE 2 |** *In ovo* **electroporation of gecko embryos. (A)** Experimental equipment. **(B)** Needle-type electrodes (CUY200S). **(C)** Two *P. pictus* eggs incubated in a small tapper with vermiculite. **(D)** Sterilization of the egg with 70% ethanol and a cotton stick. **(E)** HBSS was dropped through the hole of the shell. **(F)** The window was opened with fine forceps. **(G)** An illustration showing the position of the electrodes on the embryo. **(H)** High magnification of an electroporated embryo. Green-colored DNA solution was injected in the left lateral ventricle. **(I)** The window was sealed with a cover glass. **(J)** Incubation of operated embryos in the container.

#### *In ovo electroporation of gecko embryos*

After oviposition, the laid eggs should be isolated from the mother animal to avoid accidental crushing of the eggs. In our laboratory, the eggs are immediately transferred to a plastic container filled with dried vermiculite (**Figure 2C**). To maintain embryonic respiration, small holes are made through the lid. The egg are approximately 10 mm in diameter and 12–13 mm in length (**Figure 2C**) (Noro et al., 2009). Fertilized eggs are incubated at 28◦C in 50– 60% humidity until manipulation. *In ovo* electroporation can be performed during 10–15 d.p.o. (days of post-oviposition); after these stages, the eyes and jaws increase rapidly in size, which make it difficult to access and electroporate to brains.

To begin *in ovo* electroporation, the egg is placed on a depression slide (Matsunami, Osaka, Japan) with moistened papers (**Figure 2D**, Prowipe, Elleair, Japan). The egg is sterilized with 70% ethanol and the surface of the shell was wiped with a cotton swab (**Figure 2D**, AspureAP-7, ASONE, Japan). Pieces of vermiculites attached to the eggs are removed at this step. The position of the embryo within the egg is confirmed by illuminating the egg with a fiber optic light (SL FI-150T, Sugihara Lab Inc., Japan). To open the shell, scratch the surface of the shell with fine forceps (VIGOR TW-705#5, B. Jadbow Inc, Switzerland) under a dissecting microscope (SZ61, OLYMPUS, Japan). Because the shell of gecko eggs is extremely fragile, care should be taken to open the shell with a forceps to avoid crushing the egg. After making a small hole in the shell, 50–100µL of saline (HBSS: Hanks' buffered saline with the addition of 1% penicillin and streptomycin and 0.1% gentamycin) is added through the hole, and the further open by carefully removing the shell (**Figure 2E**). The vitelline and amniotic membranes were cut with microsurgical scissors (**Figure 2F**, RS-5620, ROBOZ, Germany). Next, 50–100µL of HBSS is further added to the egg to maintain the space for embryonic manipulation.

To prepare the DNA solution for electroporation, purified plasmid DNA vectors are dissolved in sterilized phosphate buffered saline (PBS) with a non-toxic dye (0.01% fast green). Typically, we prepare 2.5–5 µg/µL of plasmid solution for the electroporation. Holding the head of the embryo with a fine forceps, the DNA solution is injected into the left or right side of the lateral ventricle with a fine glass capillary (MODEL G-1, NARISHIGE, Japan) that is connected to a mouth-pipette (Suction tube, Drummond, USA) or miniinjector (BJ100, BEX, Japan). Subsequently, needle-type electrodes (CUY200S, NEPAGENE, Japan) is inserted into the extraembryonic space. Because DNA is negatively charged, a positive electrode was positioned at the target region (c.f., dorsal cortex), and a negative electrode was placed at the opposite side of the head (lower jaw; **Figure 2G**). The distance between the electrodes and embryos needs to be maintained (approximately 0.5–1 mm) to minimize the risk of tissue damage and hemorrhage by the direct application of electricity. Square waves of electric pulses (32 V, 50 ms, 950 ms interval, 2 or 4 pulses) are passed with an electric stimulator (SEN-3401, Nihon Kohden, Japan) or pulse generator (CUY21EDIT II, BEX, Japan). We compared survival rates of embryos at 48 h after electroporation and found that applying 4 pulses remarkably decreased the viability of gecko embryos (**Table 1**). To prevent microbe contamination, 50–100µL of HBSS with antibiotics was applied into the extra-embryonic space. After the electroporation, the shell window was sealed with a micro cover glass (**Figure 2I**, 18 mm, #1, MATSUNAMI, Japan) attached with the tissue glue (1xHistoacryl L, B.Braun, Germany). The operated eggs were kept in a sterilized moist chamber (a plastic container with respiratory holes within the lid) and incubated at 30◦C for 24 h to 1 month (**Figure 2J**).

### *In ovo electroporation of turtle embryos*

*In ovo* manipulation of turtle embryos is similar to the method for gecko embryos with slight modifications. Because the early stages of the turtle embryos are tightly attached to the inside of the shell, frequent rotation of the egg will disrupt normal development of the turtle embryos. Thus, care should be taken to maintain the orientation of the egg after oviposition (**Figure 3A**). Fertilized turtle eggs are incubated in a highly moistened container at 28◦C. We usually performed *in ovo* electroporation at stage 10–15 (10–15 days after fertilization).

**Table 1 | Efficiency of the** *in ovo* **electroporation of the gecko embryos.**

*\*Electroporation efficiency was determined by dividing the number of GFPpositive embryos by the number of collected embryos.*

*\*\*Pilot experiments for the comparison of survival rates.*

**FIGURE 3 |** *In ovo***electroporation of turtle embryos. (A)** Turtle eggs in the delivery packet. Before transferring the eggs, the top of the shell was marked to maintain an upside-down orientation. **(B)** Tools for surgical manipulation. 1: A depression slide, 2: micro scissors, 3: forceps, 4: mini drill (pinvise), 5: hand-made egg stand, and 6: a metal file. **(C)** A small scar was made on the shell with a metal file. **(D)** A pin vise was used to drill the surface of the egg. **(E)** HBSS was dropped through the small window. **(F)** The window was sealed with a cover glass after electroporation.

The position of an embryo within the egg can be monitored by illuminating the egg with a fiber light. To open the turtle eggs, a small hole is made in the shell by drilling the top of the shell with a micro drill (0.5–0.8 mm in diameter, using a pin vise, TAMIYA, Japan) under the dissecting microscope (**Figures 3B–D**). After opening a small hole on the shell, 50–100µl of HBSS with antibiotics was added through the hole, and the window was further widened by carefully removing the shell (**Figure 3E**). The chorion and amniotic membranes were cut with microsurgical scissors. After injecting a DNA solution (2.5–5µl of DNA and 0.1% fast green in PBS) into the lateral ventricle, electroporation is performed with a needle-type electrode (CUY200S), and square pulses (32 V, 50 ms, 950 ms interval, 2 pulses) are applied to the target region of the embryos using an electric stimulator or pulse generator. After electroporation, the shell window was sealed with tissue glue and a micro cover glass as in the case of the gecko eggs (**Figure 3F**). The operated embryos are maintained in a moistened chamber and incubated at 30◦C (**Table 2**).

#### *Ex ovo culture of reptilian embryos*

Exposing the embryos from the shell to the medium dramatically facilitates accessibility to the embryos and increases the efficiency of electroporation (Buchtova et al., 2008; Tschopp et al., 2014). To allow embryonic development in the medium after electroporation, we established an *ex ovo* culture system for the middle stages of reptilian embryos (**Figure 4** and **Table 3**). To begin *ex ovo* culture, fertilized gecko and turtle eggs are transferred to a glass evaporating dish filled with HBSS, and the shell was cracked within the medium with forceps to carefully expose the embryo from the extra-embryonic membrane (**Figure 4A**). The part of shell on the side of the yolk was kept to preserve the yolk sac (**Figure 4B**). Injection and electroporation can be performed within the evaporating glass (**Figure 4B**). After electroporation, the embryo was carefully transferred to a sterilized glass-made bottle (Ikemoto Rika, Tokyo, Japan) filled with 2 mL of HBSS with antibiotics (1% penicillin and streptomycin, 0.1% gentamycin) and cultured using the whole embryo culture system (Ikemoto Rika, Tokyo, Japan) in which oxygen is constantly supplied (95% oxygen, 5% carbon dioxide, 50 mL/min) to the embryos (**Figures 4C–E**). The culture temperature was maintained at 30◦C to match ideal temperature for reptilian embryogenesis. Because the embryos are damaged by bottle rotation, the culture bottles were maintained in a static position during culture. The

*\*Electroporation efficiency was determined by dividing the number of GFPpositive embryos by the number of collected embryos.*

**FIGURE 4 |** *Ex ovo* **culture system for reptilian embryos. (A)** Turtle embryos were opened in HBSS. **(B)** A turtle embryo in which a DNA solution (green color) was injected into the lateral ventricle. **(C–E)** Incubation of gecko **(C)** and turtle **(D,E)** embryos in the whole embryo culture system. Electroporated embryos were cultured in glass vials filled with HBSS. Embryo containing vials were inserted into the rotator to supply oxygen continuously. To avoid crushing the embryos, the rotating wheel was not used during the culture.

**Table 3 | Efficiency of the** *ex ovo* **culture of gecko and turtle embryos.**

*\*Electroporation efficiency was determined by dividing the number of GFPpositive embryos by the number of collected embryos.*

culture medium (HBSS) was replaced 24 h after starting the culture. The embryos can be maintained for approximately 2 days in this culture system because embryonic circulation was gradually decreases after 3 days of culture.

#### **EXPRESSION VECTORS**

Expression vectors designed for mammalian cells can be used for genetic manipulation in reptilian embryos. In general, the CAG promoter (cytomegalovirus enhancer with chicken ß-actin promoter) provides higher expression of transgenes in amniotic brains, particularly in the neural stem/progenitor cells (Niwa et al., 1991). We used several expression vectors, including pCAX-AFP (a variant form of GFP, Takahashi and Osumi, 2002) and pCAGGS-RFP (Nomura et al., 2008), which express fluorescent reporter proteins under the control of the CAG promoter. Expression vectors with Cre/loxP technology are useful for the restricted expression of transgenes in spatiotemporally controlled manners. The electroporation of Cre-recombinase expression vectors at a lower concentration (1 ng/µL) decreases the recombination frequency, which allows clonal labeling of neural stem/progenitor cells (Kato et al., 2010; Gotoh et al., 2012; Nomura et al., 2013a).

# **IMMUNOHISTOCHEMSTRY**

To perform immunohistochemical analysis, embryos are fixed with standard fixative (4% paraformaldehyde in PBS) for overnight at 4◦C and immersed in 20% sucrose for cryoprotection. The samples were embedded in OCT compound (Tissuetek, SAKURA, Japan), and 14µm of cryosections are made with a cryostat (LEICA CM1850, Germany). Several commercial antibodies are potentially applicable for immunohistochemistry in gecko and turtle embryos (**Table 4**) (Moreno et al., 2010, 2012), although not all the antibodies provide a single band with naïve brain extracts (**Figure 6** and our unpublished data).

### **REPRESENTATIVE RESULTS**

The expression of exogenous genes can be monitored soon after electroporation. We collected gecko and turtle embryos at several time points after the electroporation, and examined the expression of fluorescent reporters under a fluorescent microscope. At 2–4 days after electroporation, intense GFP expression was detected in the dorsal part of the gecko and turtle telencephalon (**Figures 5A–C**). Even at 1 month after electroporation, reporter fluorescence was still maintained in the brain (**Figures 5G–I**).

At 2 days after electroporation, GFP expression was exclusively detected in mitotic neural stem/progenitor cells that were localized at the ventricular zone of the developing gecko cortex. These neural stem/progenitor cells have a radial fiber similar to the mammalian radial glial cells, but the fibers extend in a curved manner at the neuronal layer as in the case of avian cortical radial fibers (Nomura et al., 2008, 2014). At 4 days after electroporation, GFP-positive cells migrated from the ventricular zone and positioned at the marginal zone (**Figures 5C,E,F**). GFP-positive migrating neurons in the developing gecko cortex exhibited multipolar morphology, similar to intermediate progenitor cells (IPCs) in the subventricular zone (SVZ) of the mammalian neocortex (Miyata et al., 2004; Noctor et al., 2004; Englund et al., 2005). However, unlike mammalian IPCs, we could not detect mitotic activity in the GFP-labeled multipolar cells in the gecko cortex (**Figure 5C**). This result is consistent with our observation that Tbr2-positive cells in the developing gecko cortex are post-mitotic neurons (**Figure 6**) (Nomura et al., 2013a). At 1 month after electroporation, GFP-expressing cells were still detected in the medial and dorsal cortex of gecko embryos (**Figure 5G**). Notably, these GFP-positive cells exhibited reptilian-type pyramidal neurons and extended axonal fibers toward the contra-lateral side of the cortex, which constitutes the pallial commissure in reptiles (**Figures 5H,I**).

Recent studies have shown that the transition from the multipolar to bipolar shape in the migrating neurons is critical for the development of mammalian neocortex (Noctor et al., 2004; Hand et al., 2005; Heng et al., 2008; Ohtaka-Maruyama et al., 2013; Kawauchi, 2014; La Fata et al., 2014). In contrast to the mammalian neocortex, migrating neurons in the developing gecko and turtle cortex still maintained multipolar morphology at 7 days

**Table 4 | The list of antibodies for immunohistochemistry of reptilian brains.**

*\*Specificity was examined with western blot in previous reports (Moreno et al., 2010, 2012).*

**FIGURE 5 | GFP expression in the developing gecko and turtle cortex. (A)** Developing gecko embryo after electroporation. The image was captured using an iPhone4S camera through a magnifier. **(B)** Gecko embryos at 4 days after electroporation. GFP was expressed at the dorsal part of the telencephalon (arrows). **(C)** GFP expression in the cortical neural stem/progenitor cells. Mitotic GFP-positive cells were labeled with an anti-phospho histoneH3 (PH3) antibody (red arrows). A GFP-positive cell at the outside of the ventricular zone was not mitotic (white arrow). **(D)** The distribution of GFP-positive cells in the gecko cortex at 7 days after electroporation. Arrows indicate migrating neurons **(E,F)** GFP expression in the developing turtle cortex at 4 days after electroporation. Arrows indicate migrating neurons. **(G–I)** GFP expression in the gecko cortex at 1 month after electroporation. VZ, ventricular zone; NL, neuronal layer; OB, olfactory bulb; DC, dorsal cortex; DVR, dorsal ventricular ridge. Scale bars: 25µm **(C,F)**, 50µm **(E)**.

after electroporation (**Figures 5D**, **7A–L**). To quantify the orientation of leading process in migrating neurons, the angle of the longest process in each neuron relative to the ventricular surface was quantified in mouse, gecko, turtle and chicken cortex/dorsal pallium. Comparison of leading process orientation demonstrated that all migrating neurons in the mammalian cortical plate are vertically aligned: thus, all neuronal processes are directed to the pial surface. In contrast, migrating neurons in the reptilian and avian marginal zone are not tightly aligned and extend leading process in various directions (**Figures 7M–P**). Thus, the strict alignment of bipolar migrating neurons in the cortical plate is a unique characteristic in the developing mammalian cortex. However, we also confirmed that the expression of mammalian cortical plate markers, such as Tbr1, CTIP2, and SATB2, is also detected in the developing gecko cortex (**Figures 7Q–T**) (Nomura et al., 2013a), suggesting that some of the molecular characteristics of the cortical plate neurons are conserved between the mammalian and reptilian cortex.

### **DISCUSSION**

Comparative analyses of extant amniote brains are powerful approaches to understand the evolutionary processes of the mammalian neocortex and homologous structures in non-mammalian lineages (Molnar et al., 2006; Aboitiz, 2011; Medina et al., 2013). Previous histological studies revealed that the stellate morphology of migrating neurons in the developing reptilian cortex resemble migrating neurons in the early stages of mammalian neocortex (Goffinet, 1983). Based on the ontogenic analyses, Marin-Padilla hypothesized that mammalian neocortex has dual origins: the superficial and deepest neurons (layer I and IV) retain ancestral phenotypes that are reminiscent of the amphibian or reptilian cortex, whereas the later-born cortical plate neurons (layer II-V) are recently acquired during mammalian evolution (Marin-Padilla, 1971, 1978). Our *in vivo* cell tracing analyses indicated that (1) multipolar neurons in the reptilian cortex do not exhibit mitotic activity and (2) multipolar-to-bipolar transition of migratory modes is not detected during the reptilian corticogenesis. These data support the idea that both amplification of IPCs (Martinez-Cerdeno et al., 2006; Cheung et al., 2007; Charvet et al., 2009; Puzzolo and Mallamaci, 2010) and unipolar cortical plate neurons with a "locomotive mode" are derived developmental processes in the mammalian neocortex (Aboitiz et al., 2001), through which the expansion of neuron numbers and multiple laminar structures evolved. However, the morphological similarities of migrating neurons are not always associated with common cellular dynamics and gene expression patterns. Thus, the reptilian neurons are not simply equivalent to the early stages of mammalian cortical neurons or ancestral neuronal subtypes.

In addition to cell tracing of migrating neurons, we applied several developmental techniques to analyze reptilian corticogenesis, such as (1) lineage tracing of neural stem/progenitor cells, (2) quantification of reporter activities for signaling molecules, and (3) gain- and loss-of-function analyses of specific genes in the developing reptilian cortex (Nomura et al., 2013a). These experimental approaches unveiled further unique characteristics of reptilian neural stem/progenitor cells. For example, the rates of proliferation and differentiation of reptilian cortical progenitors are very slow and contribute to the production of a lower number of cortical neurons. Some of these characteristics depend on Notch signaling, and experimental manipulation of a Notch downstream effector dramatically increased neuronal production in geckoes. We hypothesized that after the diversification of mammalian and non-mammalian amniote lineages, some critical changes in neural stem cell regulation might have occurred in the ancestral mammals and thus provided the expansion of cortical areas and massive generation of excitatory neurons (Nomura et al., 2013a,b, 2014).

Recently, whole genome sequences of Chinese softshell turtle and sew turtle have been performed, which have confirmed that turtles must be positioned phylogenetically in archosaur groups in amniotes (Wang et al., 2013). The data also demonstrated that conserved and derived genetic programs in turtle

embryogenesis contributed to the evolution of the turtle-specific body plan (Wang et al., 2013). Although genome analyses of Madagascar ground gecko have not been accomplished, draft genomes of green anole lizard (*Anolis carolinensis*), a related species to the gecko, have been published (Alfoldi et al., 2011). Genomic information of the Anolis lizard revealed unique characteristics in its genomic composition, such as homogenization of the GC content and higher number of mobile elements than other amniotes (Alfoldi et al., 2011). Additional studies of the comparative genomics of reptiles will clarify how genetic and epigenetic changes contributed to brain evolution in distinct lineages of amniotes. Genomic sequences of the Chinese softshell turtle and Anolis lizard are available at the website of the Ensemble Genome Browser (http://asia*.*ensembl*.*org/index*.*html).

Currently, we have only successfully performed *in ovo* electroporation during a narrow window of time (d.p.o. 10–16 for gecko embryos and stages 10–15 for turtle embryos). Because reptilian eyes and jaws rapidly increase in size during embryogenesis, positioning the electrodes to target dorsal cortex is technically difficult at later embryonic stages. Application of an *ex ovo* culture system for gecko and turtle embryos is also limited for 3–4 days, most likely due to the lack of some essential nutrients and/or sufficient oxygen supply. Further improvements of gene delivery tools and/or culture conditions are required to manipulate embryos at any developmental stage.

Electroporation with a transposon-mediated genomic integration system provides permanent lineage tracing in mammalian and non-mammalian vertebrates (Garcia-Moreno et al., 2014; Loulier et al., 2014). Furthermore, recent advances in genome editing tools, such as TALEN (transcription activator-like effector nuclease) and CRISPR/Cas (clustered regulatory interspaced short palindromic repeats/CRISPR-associated proteins), extend the possibility of genetic manipulation in a variety of organisms (Aida et al., 2014; Kaneko et al., 2014; Pal et al., 2014). The *in vivo* delivery of CRISPR/Cas vectors induces direct somatic recombination in target tissues, which enables the site-specific mutation of endogenous genes (Xue et al., 2014; Yin et al., 2014). The application of these new research strategies to the study of comparative brain development provides a new avenue for the understanding of the origin and evolution of amniote brains, particularly the mammalian cerebral cortex.

#### **ACKNOWLEDGMENTS**

We thank Ms Misato Kawami and Mr Kazuhiro Arimura for technical support and Happyball Python for a photograph of *Paroedura pictus*. This work was supported by a Grant-in-Aid for Challenging Exploratory Research (#24657158), Scientific Research on Innovative Areas (The Empathetic Systems, #26118510) and PRESTO, JST.

#### **REFERENCES**

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

*Received: 12 November 2014; accepted: 02 February 2015; published online: 24 February 2015.*

*Citation: Nomura T, Yamashita W, Gotoh H and Ono K (2015) Genetic manipulation of reptilian embryos: toward an understanding of cortical development and evolution. Front. Neurosci. 9:45. doi: 10.3389/fnins.2015.00045*

*This article was submitted to Neurogenesis, a section of the journal Frontiers in Neuroscience.*

*Copyright © 2015 Nomura, Yamashita, Gotoh and Ono. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.*