Generating the correct number of cortical neurons of the correct types involves changes in both the mode of division of cortical progenitors and length of their cell cycle. As discussed above, cortical progenitors switch progressively from proliferative symmetrical divisions to differentiative asymmetrical divisions during corticogenesis. At the same time, the length of the cell cycle in AP cells in mouse cortex increases from 8 h at the onset of neurogenesis (E10) to 18 h at the end (E18), largely due to lengthening of the G1 phase (Takahashi et al., 1993, 1995; Estivill-Torrus et al., 2002). Subsequent studies showed that G1 lengthening acts to promote the genesis of BP cells which have much longer cell cycle length (around 26 h in E14.5 mouse cortex) due to increase in the G1 length (Arai et al., 2011). In the macaque, AP cell cycle length is much longer than in rodents and it follows a different developmental course (Kornack and Rakic, 1998). It lengthens linearly from 23 h at E40 to 54 h at E60, then shortens linearly to 27 h by E80 (Kornack and Rakic, 1998). As the neurogenic period is 10 times longer than in rodents, macaque cortical progenitors

undergo many more rounds of division even though the duration of the cell cycle is longer (Kornack and Rakic, 1998), generating a greater number of cortical cells and a larger cortex.

There is mounting evidence that TFs exert region-specific control of cortical progenitor cell cycle progression. Pax6 is one of a number of TFs that are expressed in distinct gradients across cortical areas (Zaki et al., 2003; Sansom and Livesey, 2009; Salomoni and Calegari, 2010; Georgala et al., 2011a) and several studies have implicated Pax6 in the temporal and spatial control of cell cycle duration in cortical progenitors. Loss of Pax6 during early cortical development (E12.5) in mice *in vivo* led to a shortening of the cell cycle of progenitors coupled with higher proportions of asymmetrical divisions, resulting in a temporary increase in the production of post-mitotic neurons (Warren et al., 1999; Estivill-Torrus et al., 2002; Walcher et al., 2013). Experiments with cultured *Pax6*−*/*− mutant cortical cells showed that they too exhibit accelerated proliferation, indicating that Pax6's effects on the cell cycle are cell autonomous (Estivill-Torrus et al., 2002). In contrast, at the mid-corticogenesis stage (E15.5) *Pax6*−*/*− progenitor cells proliferated more slowly than wild type controls, showing that the effects of Pax6 on the cell cycle of cortical progenitors are context-dependent.

Gain of function studies also support the idea that Pax6 controls cell cycle progression. For example, forced expression of Pax6 impairs progenitor cell proliferation *in vitro* (Heins et al., 2002; Hack et al., 2004; Cartier et al., 2006). In *PAX77* mice, which express PAX6 protein in its normal pattern but at approximately double the wild type level, there is a reduction in the number of proliferating progenitors in the rostral and central cortex, the areas where levels of Pax6 expression are normally highest (Manuel et al., 2007). Further, APs located in rostral cortical areas proliferate more slowly in these mice (Georgala et al., 2011a,b). Thus, both gain and loss of Pax6 function studies have revealed powerful context-specific effects of Pax6 on the cell cycle of progenitor cells in a variety of cortical areas and stages.

Early in corticogenesis, expression levels of Pax6 vary between different cortical regions but its expression levels become increasingly uniform with embryonic age (Mansouri et al., 1994; Stoykova and Gruss, 1994; Manuel et al., 2007). Some of the evidence cited above indicates that the actions of Pax6 on the cell cycle of cortical progenitors are associated with its expression levels. This assumption is strongly supported by our recent study in which cell cycle parameters were systematically examined in different cortical regions at different developmental stages using mouse models with either constitutive or conditional loss of Pax6 function (Mi et al., 2013a). During early corticogenesis (E12.5), when the Pax6 gradient is steepest, areas of highest expression correlate with regions where the cell cycle duration is longest (**Figure 6**). Loss of Pax6 causes shortening of the cell cycle only in these areas (**Figure 6**). In normal embryos at older ages, the Pax6 gradient becomes progressively more uniform across the cortex, as does cell cycle length, and loss of Pax6 causes shortening of the cell cycle in all areas (**Figure 6**; Mi et al., 2013a). Taken together, these studies indicate that Pax6 primarily exerts a repressive action on the cell cycle progression of cortical progenitors, and distinct expression levels of Pax6 confer

embryos. At E12.5 Pax6 is normally expressed in a high rostral to low caudal

and caudally while the S-phase length remains unaffected.

its region and age -specific role in regulating progenitor cell proliferation.

As well as regulating the length of the cell cycle in cortical progenitors, Pax6 is involved in the control of cell cycle exit. In the *Pax6sey/sey* mutant cortex, the number of progenitor cells that exit the cell cycle during early corticogenesis (E12.5) is increased, leading to a smaller progenitor pool and an increased proportion of newborn neurons (Estivill-Torrus et al., 2002; Quinn et al., 2007). A similar effect has been seen in the eye and the spinal cord, where loss of Pax6 caused precocious neuronal differentiation due to premature cell cycle exit (Philips et al., 2005; Bel-Vialar et al., 2007). Interestingly, a study using *PAX77* mice revealed that increasing Pax6 levels also causes an increase in the proportion of progenitor cells exiting the cell cycle during late corticogenesis (E15.5; Georgala et al., 2011b), indicating that there is no simple dosage-dependent relationship between Pax6 level and progenitor cell cycle exit rate. Recently, we found that loss of Pax6 led to fewer progenitors exiting the cell cycle at the PSPB (pallial–subpallial boundary) at E12.5, the area of the telencephalon where Pax6 expression is highest, contrasting with the previous finding that *Pax6*−*/*− cortical progenitors show increased cell cycle exit at E12.5 (Estivill-Torrus et al., 2002). It appears that, although a general function of Pax6 is to influence cell cycle exit, the exact nature of its effect depends on the molecular and cellular context within which it acts. This context may change corresponding to diverse Pax6 levels in different regions at different developmental stages. In support of this assumption, a number of independent analyses demonstrated that Pax6 exerts different control functions on progenitor proliferation in different cellular contexts: Pax6 enhances proliferation in the neural retina of vertebrates (Marquardt et al., 2001) and *Drosophila* (Dominguez et al., 2004), whereas it inhibits proliferation of human glioblastoma cells (Zhou et al., 2005), cultured corneal epithelial cells (Ouyang et al., 2006) and cortical progenitors in primary cell cultures (Heins et al., 2002; Haubst et al., 2004) as well as *in vivo* (Estivill-Torrus et al., 2002; Berger et al., 2007). Therefore, it is difficult to conclude a simple universally consistent role of Pax6 in the regulation of cell cycle exit.

# Pax6 Direct Targets Regulating Progenitor Cell Proliferation

As described above, Pax6 is important for the regulation of cortical progenitor proliferation (Estivill-Torrus et al., 2002; Manuel et al., 2007; Quinn et al., 2007; Georgala et al., 2011b; Mi et al., 2013a,b). Cell cycle progression is driven by the concerted action of cyclin-dependent kinases (Cdks) and their activating partners, cyclins (Malumbres and Barbacid, 2005). Low levels of Cdk activity are sufficient for cells to transit from G1 into S phase, whereas high-levels of Cdk activity are required for the cell to undergo mitosis (Dehay and Kennedy, 2007; Kaldis and Richardson, 2012). During G1, cells integrate and respond to extracellular cues that either allow the cell to continue cycling or promote its withdrawal from the cell cycle and begin differentiation (Zetterberg et al., 1995). Progression through G1 is mainly driven by the activation of Cdk4/6 and their activating partners, D-type cyclins (Cyclin D1 and D2) as well as Cdk2 and its partner Cyclin E (Malumbres and Barbacid, 2001, 2005). Cdk activity is negatively regulated by members of the cyclin-dependent kinase inhibitors family (CKIs; Polyak et al., 1994; Sherr and Roberts, 1995; Vidal and Koff, 2000). The primary substrate of Cdks during G1 phase is the retinoblastoma protein (pRb). pRb acts as a repressor of E2F TFs through direct interaction. In quiescent cells, the activities of E2Fs are repressed through direct interaction with hypophosphorylated pRb. In cycling cells, mitogenic stimuli that elevate Cyclin levels promote pRb phosphorylation by Cyclin/Cdk complexes, which in turn prevents pRb from binding to E2Fs. Consequently, pRb/E2F complexes are dissociated and free E2Fs are now active and ready to drive the transcription of genes encoding proteins that are involved in G1 phase progression and DNA replication. Gene expression profiling studies showed that Pax6 is capable of regulating the expression of a number of cell cycle genes involved in G1 to S phase transition, such as *Cdk4, Cdk6, Ccnd1, Ccnd2, p27kip1, Cdc6, Mcm3, Mcm5, Mcm6, Cdca2* and *Cdca7*, with many of them being repressed by Pax6 (Sansom et al., 2009; Mi et al., 2013a,b). These findings are in line with the notion that Pax6 mainly acts as an important regulator of progenitor cell proliferation during early cortical development as reviewed above. Among the cell cycle genes whose expression is regulated by Pax6, Pax6 direct targets identified to date include *Cdk4*, *Cdk6*, *Mcm3*, *Cdca2*, *Cdca7,* and p27*kip1* (**Table 1**; Duparc et al., 2007; Sansom et al., 2009; Mi et al., 2013a,b).

It is interesting to note that in addition to its repressive effect, Pax6 can also activate expression of some cell cycle genes (Holm et al., 2007; Osumi et al., 2008; Sansom et al., 2009; Mi et al., 2013a,b; Xie et al., 2013). For example, Pax6 directly promotes the expression of *Cdk4* in neural stem cells in early cortical development (Sansom et al., 2009). This supports the concept that Pax6 has dual roles to both promote and inhibit cell proliferation in the developing cortex, possibly through the divergent functions of its DNA-binding subdomains (see below; Walcher et al., 2013). Using transcriptome analysis of the cortex in embryos with either gain and loss of Pax6 function, Sansom et al. (2009) showed that Pax6 regulates a core transcriptional network that controls cortical neural stem cell proliferation in a dosage-dependent manner. They found that under normal conditions *in vivo*, Pax6 has the potential to both promote and inhibit stem cell proliferation by exhibiting opposing effects on the expression of different sets of cell cycle genes, but when Pax6 levels are increased, as in transgenic cortex overexpressing Pax6, the neurogenic functions of Pax6 are dominant over its ability to promote proliferation. This study is in line with our finding that at the onset of corticogenesis, the repressive effect of Pax6 on progenitor cell proliferation and the Cdk/Cyclin mediated phosphorylation of pRb is localized to regions of cortex where Pax6 expression is normally highest, indicating a relationship between the level of Pax6 expression and its antiproliferative effect (Mi et al., 2013a). Given that Pax6 is expressed in a gradient during early cortical development, the dosagedependent effects of Pax6 on cell proliferation are important in the context of understanding how the cerebral cortex becomes divided into regions with specific cytoarchitectures and functions.

# Pax6 is an Intrinsic Determinant of Neuronal Fate in the Developing Cortex

The observations that Pax6 is specifically expressed by radial glial progenitor cells in the developing cortex and is required for them to acquire their normal morphology, gene expression pattern and cell cycle characteristics (Gotz et al., 1998), suggested that Pax6 may control their ability to generate neurons. To address this question, Heins et al. (2002) used a combination of loss and gain of function approaches. They found that cultured RGCs isolated from the developing cortex of *Pax6*−*/*− mutant mouse embryos generated half as many pure neuronal clones as RGCs from control embryos. Using a tau-GFP transgene to label and quantify neurons by fluorescence activated cell sorting (FACS), they showed that the number of neurons in the cortex of *Pax6*−*/*− mutant mouse embryos was reduced by half at E14.5 and by a third at E16.5 compared to controls, consistent with the previously reported thinner CP in *Pax6*−*/*− cortex (Schmahl et al., 1993). Forced expression of Pax6 in cells dissociated from E14 cortex or in astrocytes from postnatal cortex, via transduction of a retroviral vector containing full length Pax6 cDNA, resulted in a significant increase in the number of pure neuronal clones generated, demonstrating that Pax6 is an intrinsic determinant of neuronal fate in murine cortical progenitors (Heins et al., 2002). Mo and Zecevic (2008) showed that siRNAmediated knockdown of Pax6 expression in cultured human fetal RGCs led to a significant decrease in the number of neurons produced, suggesting that the role of Pax6 in cortical neurogenesis is maintained from rodents to humans (Mo and Zecevic, 2008).

TABLE 1 | Genes that have been identified as direct Pax6 targets in the mouse developing cortex grouped according to the developmental process that they affect.

# Pax6 Confers both Regional and Neurotransmitter Fates

The mammalian CP comprises mainly glutamatergic neurons born in the cortical VZ and GABAergic interneurons (INs). In mouse, most INs are born in the medial ganglionic eminence (MGE; ∼70%) with others coming from the caudal ganglionic eminence (CGE) and areas including the preoptic area (POA; Anderson et al., 2001; Wonders and Anderson, 2006; Fogarty et al., 2007; Gelman et al., 2009; Miyoshi et al., 2010). INs born in the LGE populate the olfactory bulb (OB) and the ventral telencephalon. Most cortical glutamatergic neurons and GABAergic interneurons are generated between E12.5 and E16.5. By E18.5, GABAergic INs are distributed throughout the cortex of wild type embryos. In contrast, the cortex of *Pax6*−*/*− mutant embryos contains large subpial ectopias composed of GABAergic INs (Kroll and O'Leary, 2005). These ectopias develop from E15.5 onward and express markers of LGE-derived INs, including Sp8, rather than markers of MGE-born INs, such as Lhx6. Their development parallels a progressive ventralization of the dorsal telencephalic VZ in *Pax6*−*/*− mutant embryos. At the start of neurogenesis (E12.5–E13.5) the most ventrolateral part of the *Pax6*−*/*− mutant cortex ectopically expresses the ventral telencephalic markers Ascl1 and Gsh2. As neurogenesis progresses ventral telencephalic identities extend dorsally such that by E14.5 most of the dorsal telencephalic VZ exhibits ventral characteristics. This progressive ventralization is accompanied by a dorsal shift in expression of markers of GABAergic INs such as Dlx1/2 and GAD67. Importantly, the INs that form the ectopias are derived from Emx1-expressing cortical progenitors which normally generate glutamatergic neurons, demonstrating that in *Pax6*−*/*− mutant mice, cortical progenitors are re-specified to generate GABAergic INs instead of glutamatergic neurons (Kroll and O'Leary, 2005).

Dlx1/2, Ascl1, and Gsh2 have been shown to promote GABAergic fate, such as the expression of Gad67 (Long et al., 2009a,b; Wang et al., 2013). Thus it is possible that Pax6 normally represses GABAergic identities in the cortex by blocking the expression of these TFs.

In *Pax6*−*/*− embryos, early born CP neurons are correctly specified as dorsal telencephalic neurons, expressing dorsal markers Math2 and Tbr1 (Schuurmans et al., 2004). They fail, however, to adopt a correct glutamatergic phenotype. Interestingly, these early born neurons are not GABAergic either (Schuurmans et al., 2004). Thus, during early corticogenesis, Pax6 participates in the specification of a correct glutamatergic phenotype but is not required to suppress GABAergic identities. Schuurmans et al. (2004) analyzed double mutants for Ngn1 and Ngn2, and showed that, during early stages of corticogenesis (E13.5), neurogenins are required to specify a glutamatergic, cortical phenotype, and repress GABAergic identities. Conversely, cortical neurons born at later stages of corticogenesis (E15.5) acquire a normal cortical and glutamatergic phenotype in *Ngn1*−*/*−*; Ngn2*−*/*− double mutants whereas in *Pax6*−*/*− embryos and in mutants lacking the orphan nuclear receptor Tlx, they express reduced levels of cortical and glutamatergic markers, compared to controls, and instead acquire a subcortical, GABAergic phenotype (Schuurmans et al., 2004). This study shows that distinct genetic programs operate during early and late corticogenesis to specify cortical regional identity and neurotransmitter fate: neurogenins are required during early stages while Pax6 and Tlx are required during later stages of corticogenesis (Schuurmans et al., 2004).

# Pax6 Confers Laminar Fate

Throughout corticogenesis, newborn neurons migrate to the CP in an inside-out sequence such that deep layers are formed first and superficial layers form later. The final laminar position and subtype of cortical projection neurons is dependent on the time at which progenitors exit the cell cycle. In *Pax6*−*/*− mutant embryos, the SVZ and superficial cortical layers are defective, as shown by absence of expression of Svet1 (which marks the SVZ and superficial layers), while the VZ and deep CP layers are unaffected, as shown by the expression of Otx1 (Tarabykin et al., 2001).

Examination of cortical lamination in *Ngn1*−*/*−; *Ngn2*−*/*− mutants showed that the expression of markers of deep cortical layers is reduced (Tbr1, Er81) while markers of superficial layers are expressed normally, indicating that late-born cortical neurons are correctly specified (Schuurmans et al., 2004). In contrast, no defects are detected in the deeper cortical layers of *Pax6*−*/*−, *Tlx*−*/*−, or *Pax6*−*/*−; *Tlx*−*/*− double mutants. Markers of upper layers, however, are downregulated in Pax6 and Tlx single mutants and completely lost in *Pax6*−*/*−;*Tlx*−*/*− embryos suggesting that Pax6 and Tlx cooperate to specify the identity of late-born cortical neurons (Schuurmans et al., 2004).

Study of cortical lamination in *Pax6*−*/*− mutants is limited by the fact that they die perinatally while cortical neurons do not reach their final location until around postnatal day 8 (P8). Therefore, more recent studies on the role of Pax6 in cortical lamination have used conditional mutants in which *Pax6* is inactivated specifically in the cortex and at later stages of cortical development and which survive postnatally. When *Pax6* was deleted specifically in the cortex from the onset of corticogenesis, markers of superficial layers were either downregulated or lost completely (Tuoc et al., 2009). In contrast, when *Pax6* was deleted in RGCs after the generation of deep cortical layers (between E14.5 and E16.5) using an hGFAPcre transgenic line, the specification and number of late born neurons was unaffected (Tuoc et al., 2009), leading to the conclusion that Pax6 is not required for the specification of late born cortical neurons and that the defects of superficial layers observed in *Pax6*−*/*− and *Pax6*cKO mutants are a result of earlier defects in RGC proliferation. In a more recent study, Georgala et al. (2011b) used an *Emx1-creErT2* transgenic line to remove Pax6 specifically in the cortex between E12.5 and E13.5, before superficial layer neurons are born. The resulting cKO mutant brains showed a dramatic reduction in the thickness of the superficial cortical layers, compared to control embryos. Late-born neurons were birth-dated by injecting BrdU at E15.5 and stained for expression of Cux1, a marker of upper layer (II to IV) pyramidal neurons. There was a substantial reduction in the number of E15.5 born neurons in the superficial cortical layers of the *Pax6-cKO* mutants and many fewer late-born neurons expressed Cux1 (Georgala et al., 2011b), indicating that Pax6 plays a crucial role in generation and specification of late born cortical neurons. Note that Pax6 protein was lost slightly earlier in the conditional mutants generated by Georgala et al. (2011b) than in those used by Tuoc et al. (2009), potentially explaining the difference in outcomes between the two studies.

# Direct Transcriptional Targets of Pax6 Regulate Basal Progenitor Genesis, Neurogenesis, and Cell Fate Specification

Pax6 exerts many of its effects, at least in part, by directly regulating the expression of specific target genes (**Table 1**). As described above, work in the mouse has shown that Pax6 expressing APs that give rise to BPs transiently express the proneural TF Neurogenin 2 (Ngn2). BPs do not express Pax6 but do express the TF Tbr2. Loss of Pax6 function results in loss of Ngn2 expression in the cortical VZ and downregulation of Tbr2 in the SVZ (Stoykova et al., 2000; Toresson et al., 2000; Yun et al., 2001; Scardigli et al., 2003; Holm et al., 2007; Quinn et al., 2007; Sansom and Livesey, 2009), suggesting that the development of BPs is impaired. Conversely, Pax6 overexpression results in a marked increase in both Ngn2 and Tbr2 expression (Scardigli et al., 2003; Sansom and Livesey, 2009), each of which have been shown to be directly regulated by Pax6 (Scardigli et al., 2003; Sansom and Livesey, 2009; Tuoc et al., 2013a). Scardigli et al. (2003) showed that Pax6 directly binds and regulates the activity of the E1 enhancer of *Ngn2*, which regulates *Ngn2* expression in the cortex and the ventral spinal cord.

Another direct target that is positively regulated by Pax6 is the anti-neurogenic transcriptional co-repressor Tle1, a member of the Groucho/transducin-like enhancer of split (Gro/Tle) family (Sansom et al., 2009; Tuoc et al., 2013a). Gro/Tle proteins are expressed in proliferating neural progenitor cells where they promote maintenance of the undifferentiated state by inhibiting/delaying neuronal differentiation (Buscarlet and Stifani, 2007; Jennings and Ish-Horowicz, 2008).

Pax6 also directly inhibits the expression of Olig2 (Jang and Goldman, 2011), a TF critical for glial cell fate determination (Marshall et al., 2005). Olig2 is expressed in a subset of cells in the rat neonatal SVZ, which differentiate into glia. Forced expression of Pax6 in these Olig2+ cells, using a retrovirus, caused downregulation of Olig2 and a shift toward neuronal fate (Jang and Goldman, 2011). These authors showed that Pax6 can directly bind to Olig2's promoter and repress its activity.

Pax6 directly regulates markers of specific neuronal subtypes including Er81 and Cux1 (Tuoc and Stoykova, 2008a; Sansom et al., 2009; Tuoc et al., 2013a). Er81 is a TF expressed in cortical progenitors, in a rostrolateral-high to caudomedial-low gradient, and in a subset of pyramidal cells scattered through layer 5 (Yoneshima et al., 2006; Tuoc and Stoykova, 2008a). In the absence of Pax6, cortical progenitors do not express Er81 and the rostrolateral cortex lacks Er81 positive layer 5 neurons. Cux1 is a marker of upper layer (II to IV) pyramidal neurons and functions as a negative regulator of dendritic complexity for these neurons (Cubelos et al., 2010, 2014; Li et al., 2010). As mentioned above, loss of Pax6 results in a dramatic reduction of Cux1+ upper layer neurons, suggesting that the specification of late born neurons is impaired (Georgala et al., 2011b).

How does a single gene exert such a wide range of biological effects? Part of the answer to this question is suggested by the findings that the Pax6 protein contains two distinct DNAbinding domains, one of which is regulated by alternate splicing and that Pax6 is able to interact with a number of co-factors, which influence its activity. This issue is discussed in the next section.

# Complexity of Pax6 DNA-Binding Properties and its Distinct Functions

Pax6 binds to target DNA sequences through one or both of two DNA-binding domains, the paired domain (PD) and the homeodomain (HD; Bertuccioli et al., 1996; Jun and Desplan, 1996; Sheng et al., 1997; Singh et al., 2000). Transcriptional regulation of Pax6 target genes is mediated by a carboxy terminal proline/serine/threonine (PST) rich transactivation domain (Singh et al., 1998, 2001; Tang et al., 1998; **Figure 7**).

The PD is an evolutionarily conserved 128 amino acid DNA-binding domain shared by all members of the paired box (Pax) family of TFs (Pax1-Pax9; Chalepakis et al., 1992; Gruss and Walther, 1992). The Pax6 PD can be structurally and functionally separated into two independent DNA-binding globular helix-turn-helix subdomains, PAI and RED (Puschel et al., 1992; Czerny et al., 1993; Jun and Desplan, 1996; **Figure 7A**). The well-established DNA-binding motif for canonical Pax6 PD, known as P6CON, is a bipartite 20 base pair DNA sequence whose 5 -half is recognized by the PAI subdomain and 3 -half by the RED subdomain (Czerny et al., 1993; Epstein et al., 1994; **Figure 7A**). The crystal structure of the Pax6 PD – P6CON complex revealed direct interactions between both PAI and RED subdomains, the linker regions of PAI and RED subdomains and the N-terminal β-turn subdomain with individual bases and phosphate residues of cognate DNA (Czerny et al., 1993; Epstein et al., 1994; Xu et al., 1999a). Pax6 also contains an internal, paired-type HD (Czerny et al., 1999; Eberhard and Busslinger, 1999) which binds DNA in the form of a homodimer recognizing a symmetric binding site (Wilson et al., 1993; Eberhard and Busslinger, 1999; Mikkola et al., 2001; **Figure 7B**).

Although different Pax6 DNA-binding domains can independently bind to their "optimal" binding sites, using individual consensus binding motif to predict and confirm candidate Pax6 binding sites has proven to be very difficult (Shimoda et al., 2002; Visel et al., 2007; Wolf et al., 2009). Unlike sequence specific DNA-binding proteins which harbor only one DNAbinding domain, Pax6 exhibits increased DNA-binding complexity because of its multiple DNA-binding subdomains (PAI, RED, and HD) which can be used in varying combinations when binding to DNA (Cvekl et al., 2004; Grapp et al., 2009). A recent study further reinforced the complexity of Pax6 DNA-binding properties by systematically examining coordinated actions of different Pax6 binding subdomains and identified a number of novel variants of Pax6 DNA-binding motifs which can be bound by different combinations of Pax6 binding subdomains (Xie and Cvekl, 2011). As Pax6 can function either as a transcriptional activator or repressor (Wolf et al., 2009), it is possible that some of these binding motif variants may induce structural changes in Pax6 proteins that enable them to switch their function between transcriptional activation and repression.

The complexity of the DNA binding properties of different Pax6 subdomains implies that the subregions exert distinct functions during embryonic cortical development. Pax6 mutant mouse lines with disruption of either the PD or the HD have been used to investigate the impact of different Pax6 binding domains on telencephalic development. Haubst et al. (2004) showed that the PD alone is necessary and sufficient for the regulation of most aspects of telencephalic development including neurogenesis, progenitor cell proliferation, and patterning in the developing cortex. In contrast, HD-specific mutations affected only subtle aspects of telencephalic development, such as establishment of the PSPB, and had no obvious effect on neurogenesis or proliferation (Haubst et al., 2004). This suggests that the majority of Pax6's effects during telencephalic development are mediated by the PD with potentially different contributions by its subdomains. A recent study further dissected the roles of the different

subdomains of the Pax6 PD during telencephalic development by examining mice with point mutations in its individual subdomains PAI and RED (Walcher et al., 2013). This showed that both PAI and RED act largely in a redundant manner in patterning of the telencephalon, with only the PAI mutation resulting in the ectopic expression of subpallial genes such as *Gsx2* and *Olig2* into the pallium, suggesting that RED subdomain on its own is not sufficient to regulate telencephalic patterning. In addition, neurogenesis was affected only by the PAI subdomain mutation, phenocopying the neurogenic defects observed in full *Pax6*−*/*− mutants. Strikingly, this study also revealed that subdomains of Pax6 PD have distinct roles in regulating cortical progenitor cell proliferation, with mutations affecting the PAI and RED subdomains respectively reducing and increasing the number of mitoses.

# Alternatively Spliced Isoforms of Pax6 and their Functions

In addition to Pax6 and Pax6(5a) a number of other splice variants of Pax6 have been reported, named p43, p33 and p32, each of which is less abundant and mostly localized in the cytoplasm (Carriere et al., 1993; Jaworski et al., 1997; Pinson et al., 2006). The 14 amino acid insertion in Pax6(5a) destroys the DNA-binding capacity of the PAI subdomain, leaving the RED subdomain of the PD and the intact HD for DNA binding (Epstein et al., 1994; Kozmik et al., 1997). The PD in Pax6(5a) binds to a distinct DNA sequence from that bound by the canonical Pax6 PD, that has been named 5aCON (Epstein et al., 1994). Direct comparison of the DNA binding sites recognized by Pax6 PD and PD(5a) showed partial sequence homology between the 3' halves of P6CON and 5aCON (Epstein et al., 1994; Duncan et al., 1998, 2000). This homology allows Pax6 (which contains the canonical PD) to bind to both P6CON and 5aCON sites, whereas Pax6(5a) can only interact with the 5aCON site (Epstein et al., 1994).

The majority of published molecular studies have focused on Pax6, as it contains the canonical PD and appears to be more abundant than Pax6(5a) (Carriere et al., 1993; Richardson et al., 1995; Koroma et al., 1997). During early development, *Pax6* and *Pax6(5a)* transcripts are expressed in a ratio of 8:1 in the mouse forebrain, falling to 3:1 between E12.5 and E14.5 (Kozmik et al., 1997; Pinson et al., 2006). A reduced ratio between Pax6 and Pax6(5a) was also seen during chick retina development (Azuma et al., 2005).  These findings raise the possibility that expression of the two main Pax6 isoforms are regulated in a tissue- and temporal-specific manner, thereby affecting which target genes they recognize.

Although Pax6 and Pax6(5a) are commonly co-expressed, there is evidence that they exert distinct functions during embryonic development. For example, overexpression of either Pax6 or Pax6(5a) in chick retina showed that both isoforms led to increased retinal progenitor cell proliferation, but Pax6(5a) induced ectopic differentiation of the retina to a stronger degree than Pax6 (Azuma et al., 2005). Similarly, Pax6(5a) strongly induces neural differentiation of murine embryonic stem cells *in vitro*, while Pax6 does not have such a strong effect (Shimizu et al., 2009). In the embryonic mouse forebrain, the different roles of the two Pax6 isoforms in regulation of cell proliferation and differentiation were also established from studies of Pax6 mutants. It was shown that mutations that abolish the binding property of the PD affect both proliferation and differentiation, while a PD(5a) mutation only affects proliferation and has no effect on cell fate/differentiation in mouse developing telencephalon (Haubst et al., 2004). In a more recent study, Pax6(5a) specific target genes were identified through microarray analyses of gene expression profiles in cell lines stably expressing either Pax6 or Pax6(5a) (Kiselev et al., 2012). This revealed that a number of genes involved in cell proliferation, differentiation, and migration are differentially regulated by Pax6 and Pax6(5a), providing a potential molecular basis by which Pax6 and Pax6(5a) could exert their distinct biological functions.

# Co-Factors Influence Pax6's Direct Regulation of Target Genes Involved in Embryonic and Adult Neurogenesis

It is becoming clear that transcriptional regulation is highly dependent on the molecular and cellular context, one reason for which is that TFs do not act on their own but are dependent on interactions with other co-factors to exert their functions (Oikawa and Yamada, 2003; Westerman et al., 2003). These interactions introduce more specificity into the regulatory function of a given TF in a particular cellular context. A number cofactors that can interact with Pax6 have been identified. For example, Pax6 cooperates with Sox2 to target the lens-specific enhancer of the δ*-crystallin* gene (Kamachi et al., 2001); with MDIA to influence Pax6's functions in cerebellar granule cells (Tominaga et al., 2002); with Trim11 to mediate Pax6 degradation through the ubiquitin proteasome system (UPS) and modulate Pax6 transcriptional activity (Cooper and Hanson, 2005; Tuoc and Stoykova, 2008b). More recently, Brg1/Brm associated factors complex (BAF) and Meis2 have been identified as two new co-factors that affect Pax6's direct regulation of target genes expression, in turn regulating its role in both embryonic and adult neurogenesis (Ninkovic et al., 2013; Tuoc et al., 2013a,b; Agoston et al., 2014).

# BAF Complex

A number of chromatin remodeling factors are known to play an important role during mammalian neural development, including the BAF complex (Ho et al., 2009a,b; Wu, 2012; Yip et al., 2012; Tuoc et al., 2013a,b). Interestingly, the function of the BAF complex in regulating neural development is dependent on its subunit composition. In mammals, BAF complexes contain two ATPase subunits Brg1 or Brm (Brahma), which are mutually exclusive and essential for remodeling activity, in combination with at least 15 different BAF (Brg1/Brm-associated factor) subunits (Ho et al., 2009a; Singhal et al., 2010). Certain BAF subunits have restricted expression patterns and thus could define tissueor cell-type-specific BAF complexes. For example, the composition of BAF complexes in ES cells (esBAF) is defined by the incorporation of Brg1 but not Brm, BAF155 but not BAF170, and BAF60A but not BAF60C (**Figure 8**; Ho et al., 2009a,b). However, during the transition from ES cells to neural progenitors, the composition of BAF complexes is correspondingly changed, defining the neural progenitor specific BAF complex (npBAF; Lessard et al., 2007; Yan et al., 2008; Ho et al., 2009a). This process is accompanied by induction of BAF170 expression, replacing one of the BAF155 subunits and the recruitment of Brm as the ATP-ase subunit in the npBAF complex.

In two recent studies, Tuoc et al. (2013a,b), described an elegant model in which time-specific binding of BAF subunits including BAF170, BAF155, and Brm-ATPase to Pax6 modulates the expression of Pax6 target genes (*Tbr2, Cux1,* and *Tle1*) involved in the specification and generation of BPs and upper layer neuronal fate in the developing cortex. Interestingly, this model involves competition between BAF170 and BAF155 subunits in the BAF complex, which modulates chromatin modifications and binding of the Pax6/REST co-repressor complex to Pax6 target genes, thereby affecting their expression. During early neurogenesis (E12.5 to E14.5), the presence of BAF170 in the Pax6-BAF complex prevents the euchromatin state (a lightly packed form of chromatin) of the Pax6 target genes *Tbr2, Cux1* and *Tle1*, limiting the ability of Pax6 to bind them, thereby inhibiting their expression and promoting direct neurogenesis (RGCs directly give rise to neurons) at this developmental stage. In contrast, after E14.5 the expression of BAF170 decreases, accompanied by enhanced expression of BAF155, leading to a relaxed chromatin state of the promoters of the Pax6 target genes mentioned above, thereby enhancing their expression and in turn promoting indirect neurogenesis (RGCs indirectly give rise to neurons through generating BPs). This study provides novel insight into the interaction between BAF complex and Pax6 in the generation of BPs and the mode of neurogenesis in the developing cortex. In another recent study, Ninkovic et al. (2013) showed that Pax6 directly interacts with Brg1-ATPase of the BAF complex to modulate expression of genes involved in neuronal fate specification in adult neural progenitors. This study indicated that direct interaction of Pax6 with the Brg1/BAF complex enhances the expression of Pax6 target neurogenic TFs including *Sox11, Nfib,* and *Pou3f4* which form a cross-regulatory network driving neurogenic fate during adult neurogenesis. This study improved our understanding of Pax6-mediated adult neurogenesis and shed light on molecular mechanisms underlying the function of Pax6 in cell fate specification and conversion.

# Meis2

Meis2 belongs to the TALE (three amino acid loop extension) family of atypical HD-containing TFs. It can form heteromeric complexes with other transcriptional regulators to exert its functions during a range of biological processes (Moskow et al., 1995; Chang et al., 1997; Knoepfler et al., 1999; Moens and Selleri, 2006). Meis proteins control cell cycle progression and cell fate specification of different stem and progenitor cell types in a variety of tissues including liver, heart, retina, and brain during development (Mercader et al., 1999; Hisa et al., 2004; Bessa et al., 2008; Heine et al., 2008; Agoston and Schulte, 2009; Choe et al., 2009; Agoston et al., 2012, 2014; Paige et al., 2012). Recently the functional importance of Meis2 in adult neurogenesis has been discovered. It is involved in coordinated actions with Pax6 (Agoston et al., 2014). During mouse adult neurogenesis, Meis2 is strongly expressed in the SVZ and rostral migratory stream (RMS) and in a subset of OB interneurons (Agoston et al., 2014). Agoston et al. (2014) found that Meis2 can physically interact with Pax6 in cells in the SVZ and OB. This study also showed that the pro-neurogenic function of Pax6 in the adult SVZ requires direct interaction with Meis2. Meis2 has transcriptional activator function and Agoston et al. (2014) further confirmed that it acts as a Pax6 co-factor, binding to regulatory elements of the *Dcx* gene to promote its expression during adult SVZ neurogenesis.

# Conclusion

The TF Pax6 is one of the most intensively studied high-level developmental regulators, playing crucial roles in brain development and implicated in disease. Numerous studies, including those from our group, have shown that in both humans and mice many abnormalities resulting from mutations in *PAX6/Pax6* are in the cerebral cortex, where they arise largely because Pax6 is required for spatio-temporal control of many biological processes including cortical stem and progenitor cell proliferation and differentiation, neurogenesis, cell and laminar fate specification, neuronal migration, axon guidance and cortical arealization during early corticogenesis. However, we are a long way from having a comprehensive knowledge of exactly how Pax6 exerts such complex actions. In particular, there are at least three major challenges toward improved understanding of Pax6 functions. First, how does such highly complex expression pattern of Pax6 come about? As described above, the transcriptional regulation of *Pax6* is particularly complex and involves both short-range and long-range control mechanisms. Future studies are required to gain better knowledge of tissue- and time-specific regulatory elements responsible for spatiotemporally and quantitatively correct expression of Pax6. Secondly, how does Pax6 expression level influence its functional outputs? Our studies along with others have shown that Pax6 expression levels are crucial for its functions such as regulating progenitor cell proliferation and differentiation, and that mutations affecting Pax6 protein levels cause neurodevelopmental disorders in both humans and mice. Now it is time to ask exactly how Pax6 levels influence its actions at the molecular level. It will be of great interest to examine how varying levels of Pax6 influence its molecular actions at different cell cycle phases, which will be critical to obtain a clearer understanding of how Pax6 functions during the dynamics of cell division. Finally and probably most importantly, we are still far away from a comprehensive understanding of the complex gene networks that Pax6 regulates and how these networks change spatially and temporally during the course of cortical development. This represents a major challenge since Pax6 likely exerts direct and indirect regulation of different sets of downstream genes simultaneously, with context-dependent variation in the strength of their positive or negative effects.

# Acknowledgments

Research in the authors' laboratory was funded by the MRC (MR/J003662/1). Human embryonic and fetal material was provided by the Joint MRC/Wellcome Trust (grant # 099175/Z/12/Z) Human Developmental Biology Resource (www.hdbr.org).

# References

in the outer subventricular zone of the primate. *Neuron* 80, 442–457. doi: 10.1016/j.neuron.2013.09.032 S0896-6273(13)00863-5

cell cycle and specifies cortical cell identity by a cell autonomous mechanism. *Dev. Biol.* 302, 50–65. doi: 10.1016/j.ydbio.2006.08.035

**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 Manuel, Mi, Mason and Price. 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.*

# Real-time imaging of bHLH transcription factors reveals their dynamic control in the multipotency and fate choice of neural stem cells

#### Edited by:

Takeshi Kawauchi, Keio University School of Medicine/PRESTO, Japan

#### Reviewed by:

Noelia Urbán, The Francis Crick Institute - Mill Hill Laboratory, UK Stewart Anderson, Children's Hospital of Philadelphia, USA William A. Harris, University of Cambridge, UK

#### \*Correspondence:

Itaru Imayoshi, The Hakubi Center, Institute for Virus Research, Kyoto University, Shogoin-Kawahara 53, Sakyo-ku, Kyoto 606-8507, Japan<EMAIL_ADDRESS>Ryoichiro Kageyama, Institute for Virus Research, Kyoto University, Shogoin-Kawahara, Sakyo-ku, Kyoto 606-8507, Japan<EMAIL_ADDRESS>
> Received: 28 January 2015 Accepted: 13 July 2015 Published: 04 August 2015

#### Citation:

Imayoshi I, Ishidate F and Kageyama R (2015) Real-time imaging of bHLH transcription factors reveals their dynamic control in the multipotency and fate choice of neural stem cells. Front. Cell. Neurosci. 9:288. doi: 10.3389/fncel.2015.00288

#### Itaru Imayoshi 1, 2, 3, 4 \*, Fumiyoshi Ishidate3 and Ryoichiro Kageyama2, 3, 5 \*

1 The Hakubi Center, Kyoto University, Kyoto, Japan, 2 Laboratory of Growth Regulation, Institute for Virus Research, Kyoto University, Kyoto, Japan, 3 World Premier International Research Initiative–Institute for Integrated Cell-Material Sciences, Kyoto University, Kyoto, Japan, 4 Precursory Research for Embryonic Science and Technology, Japan Science and Technology Agency, Saitama, Japan, 5 Core Research for Evolutional Science and Technology, Japan Science and Technology Agency, Saitama, Japan

The basic-helix-loop-helix (bHLH) transcription factors Ascl1/Mash1, Hes1, and Olig2 regulate the fate choice of neurons, astrocytes, and oligodendrocytes, respectively; however, these factors are coexpressed in self-renewing multipotent neural stem cells (NSCs) even before cell fate determination. This fact raises the possibility that these fate determination factors are differentially expressed between self-renewing and differentiating NSCs with unique expression dynamics. Real-time imaging analysis utilizing fluorescent proteins is a powerful strategy for monitoring expression dynamics. Fusion with fluorescent reporters makes it possible to analyze the dynamic behavior of specific proteins in living cells. However, it is technically challenging to conduct long-term imaging of proteins, particularly those with low expression levels, because a high-sensitivity and low-noise imaging system is required, and very often bleaching of fluorescent proteins and cell toxicity by prolonged laser exposure are problematic. Furthermore, to analyze the functional roles of the dynamic expression of cellular proteins, it is essential to image reporter fusion proteins that are expressed at comparable levels to their endogenous expression. In this review, we introduce our recent reports about the dynamic control of bHLH transcription factors in multipotency and fate choice of NSCs, focusing on real-time imaging of fluorescent reporters fused with bHLH transcription factors. Our imaging results indicate that bHLH transcription factors are expressed in an oscillatory manner by NSCs, and that one of them becomes dominant during fate choice. We propose that the multipotent state of NSCs correlates with the oscillatory expression of several bHLH transcription factors, whereas the differentiated state correlates with the sustained expression of a single bHLH transcription factor.

Keywords: bHLH, neural stem cells, oscillation, fluorescent protein, imaging

# Introduction

Neural stem cells (NSCs) are multipotent and self-renewable cells that can give rise to neurons, astrocytes, and oligodendrocytes (Fishell and Kriegstein, 2003; Götz and Huttner, 2005; Kriegstein and Alvarez-Buylla, 2009). During brain development, basic helix-loop-helix (bHLH) transcription factors play pivotal roles in the self-renewal of NSCs and fate determination of neurons, astrocytes, and oligodendrocytes (Bertrand et al., 2002; Ross et al., 2003; Meijer et al., 2012; Wilkinson et al., 2013; Imayoshi and Kageyama, 2014a). bHLH transcription factors, such as Hes1 and Hes5, regulate the self-renewal of NSCs as downstream effectors of Notch signaling, whereas proneural bHLH transcription factors, such as Ascl1 and Neurog1/2, promote neuronal differentiation. Other bHLH transcription factors, e.g., Olig1 and Olig2, regulate oligodendrocyte differentiation. However, in addition to Hes1/Hes5, some bHLH fate determination factors, such as Ascl1 and Olig2, are also known to have roles in NSC maintenance or proliferation. In addition, Hes1 induces astrocyte formation at later stages. Currently, it is not understood completely how these various and sometimes opposing functions of each bHLH transcription factor in the self-renewal and fate-choice events of NSCs are achieved (Vasconcelos and Castro, 2014; Imayoshi and Kageyama, 2014a,b). We previously found that transcription of some bHLH genes is differentially controlled in NSCs and differentiating cells (Shimojo et al., 2008). Therefore, we decided to analyze the expression dynamics of bHLH transcription factors in more detail.

# Heterogeneous Expression of bHLH Transcription Factors in Cultured NSCs

NSCs can be cultured in an undifferentiated state in vitro. For instance, in the presence of epidermal growth factor and basic fibroblast growth factor, NSCs can be expanded from enzymatically dissociated brain tissues in a monolayer condition (NS cell culture) (Conti et al., 2003). NSCs can reportedly be established from brains at various stages, from embryos to adults. Furthermore, the purity of these cultures is very high; more than 99% of cells express NSC marker proteins, such as Nestin. Cultured NSCs also express bHLH transcription factors, including Hes1, Ascl1, and Olig2 (Imayoshi et al., 2013a).

In actively-dividing and self-renewing NS cells, Hes1, Ascl1, and Olig2 bHLH transcription factors are expressed (**Figure 1A**). Compared with the homogeneous expression of Sox2, which is a member of the high-mobility-group transcription factor family, the expression of all three bHLH transcription factors is variable among cells, indicating their dynamic expression (**Figure 1A**) (Imayoshi et al., 2013b). In order to elucidate the heterogeneous expression of bHLH transcription factors, we adopted a real-time imaging strategy to monitor the expression dynamics of bHLH proteins in cultured NSCs.

FIGURE 1 | Expression of bHLH factors in NS cells and fluorescent reporter-bHLH fusion constructs. (A) Self-renewing NS cells were immunostained with anti-Sox2, anti-Hes1, anti-Ascl1, and anti-Olig2 antibodies. Hes1, Ascl1, and Olig2 expression levels were variable while another NSC-specific factor, Sox2, was expressed at a relatively constant level. (B) Venus-Hes1 fusion knock-in mouse strain. (C) Venus-Ascl1 fusion BAC Tg mouse strain. (D) mCherry-Olig2 fusion BAC Tg mouse strain. Coding regions of bHLH factors are shown in gray. The pictures and graphs of this figure are reprinted from Supplementary Figures S1C,E; S2A; S6A–D of Imayoshi et al. (2013b).

# Fluorescent Reporter-fusion Knock-in or Bacterial Artificial Chromosome (BAC) Transgenic (Tg) Strategy for Monitoring Protein Expression Dynamics of bHLH Transcription Factors

Fusion reporter constructs are used to monitor the expression dynamics of proteins of interest in living cells (**Figures 1B–D**) (Miyawaki, 2011; Imayoshi et al., 2013a). In addition to fluorescent reporter proteins, bioluminescent reporters, such as luciferase, are also effective. One advantage of the reporter-fusion strategy is that the half-life and cellular localization of proteins basically depend on the fused target protein, at least in the case of bHLH transcription factors. Most bHLH transcription factors have very short half-lives, such as ∼20 min, and therefore reporter-bHLH fusion proteins are also very unstable.

In order to analyze precisely the fine expression dynamics of bHLH transcription factors, it is necessary that reporterbHLH fusion proteins are expressed under the control of their original promoter/enhancer sequences. One of the reliable ways to mimic the original regulation of bHLH gene expression is the knock-in strategy (Imayoshi et al., 2013a,b). By homologous recombination in embryonic stem cells, a reporter coding sequence is inserted into the N− or C− terminus of a bHLH gene so that the reporter-bHLH fusion protein is expressed from the original bHLH gene locus. As far as we know, the reporter-bHLH fusion protein is functional and, in most cases, the allele carrying the knocked-in fusion reporter functions as normally as the wild-type allele. Another strategy to express reporter-bHLH fusion proteins optimally is the generation of Tg mice with a BAC. Using homologous recombination in Escherichia coli, we can edit and modify BACs (Lee et al., 2001). BACs can contain very long DNA sequences, such as ∼200-kb. Therefore, by utilizing BAC clones with the promoter/enhancer sequences of a bHLH gene, it is possible to make transgene constructs designed to express reporter-bHLH fusion proteins under its original regulatory elements. From the Tg mouse founder lines carrying BAC transgene constructs, we can choose ones with a transgene copy number of only one by Southern blotting or real-time PCR analysis. These one-copy BAC Tg mice work as reporter strains for real-time imaging of the expression dynamics of bHLH transcription factors (Imayoshi et al., 2013b). One concsern regarding the reporter-fusion BAC Tg mouse strategy is that the number of functional bHLH gene alleles is increased from two to three. Ideally, reporter expression should be analyzed in a heterozygous-mutant background of the target gene by crossing with a mouse strain harboring a null allele. However, at least in the case of bHLH transcription factors, such as Hes1, Ascl1, or Olig2, the addition of the reporter-fusion transgenes dose not apparently affect the normal development of the nervous system (Imayoshi et al., 2013b).

As far as we know, the N-terminal fusion of a reporter proteins in both knock-in and BAC Tg mice shows better results than for a C-terminal fusion. The fluorescence or bioluminescence of a reporter protein is brighter when it is fused to the N-terminus than to the C-terminus of bHLH transcription factors. This is probably because translation starts from the N-terminus, so a reporter protein located at the N-terminus starts to work earlier during its expression.

For quantitative live-imaging of fluorescent reporters, we established NS cell cultures from each knock-in or BAC Tg mouse strain (Imayoshi et al., 2013b). In fluorescent protein-bHLH fusion reporter mice, Venus-Hes1 fusion knock-in, Venus-Ascl1 fusion BAC Tg, and mCherry-Olig2 fusion BAC Tg mouse strains (**Figures 1B–D**), the expression levels of fusion proteins and endogenous bHLH proteins were highly correlated to each other in cultured NSCs (Imayoshi et al., 2013b). Therefore, we conducted real-time imaging of fluorescent protein-bHLH fusion reporters in cultured NSCs with confocal fluorescence microscopy.

# Real-time Imaging of Fluorescent Reporter-fusion bHLH Proteins with Confocal Fluorescent Microscopy

For real-time imaging of fluorescent proteins with low expression levels, it is important to prepare a highly sensitive and lownoise imaging system. As we mentioned earlier, the mRNA and protein of most bHLH transcription factors have very short half-lives (e.g., ∼20 min): therefore, the reporter-bHLH fusion proteins are also very unstable (Imayoshi et al., 2013a,b). We can measure the decay time of reporter-bHLH fusion products by pharmacologically blocking the synthesis of new cellular proteins with cycloheximide. Indeed, the reporter activity of bHLH-fusion products are decayed with the same time course as the original bHLH proteins (data not shown). Furthermore, in cultured NSCs derived from knock-in or BAC Tg mice, reporterbHLH fusion proteins are expressed from a limited number of gene alleles. Therefore, the fluorescent signals of reporter-bHLH fusion proteins in NSCs are very dim and sometimes invisible by simple observation through ocular lenses.

To acquire these very weak fluorescent signals, we utilized a confocal microscope equipped with a highly sensitive detector (Imayoshi et al., 2013b). In some cases, the fluorescent signal intensity of reporter-bHLH fusion proteins is much weaker than that of the autofluorescence of cultured NSCs. To overcome this problem, a spectral imaging technique is very effective. Spectral imaging and linear unmixing has become an important tool in confocal fluorescence microscopy to discriminate between fluorescent signals with overlapping spectral characteristics. For example, when cultured NSCs are illuminated with a 514-nm argon laser, the autofluorescent signal appears at approximately 560 nm. Real signals from Venus-Hes1 or Venus-Ascl1 fusion proteins are observed approximately 530 nm, and their intensity is weaker than the autofluorescence of NSCs. However, when the fluorescent signals are acquired with a spectral GaAsP array detector, specific Venus-bHLH signals can be separated from autofluorescence by applying linear unmixing algorithms.

In addition to the highly sensitive and spectral imaging techniques, imaging systems should be optimized for lownoise and reliable quantitative analysis. As the fluorescent signal intensity of reporter-bHLH fusion proteins is very weak, contamination with noise signals impairs the reliable quantitative analysis of the expression dynamics of bHLH transcription factors. To prevent bleaching the fluorescent reporters and avoid photodamage to NSCs by the excitation lasers, the sensitivity of the imaging system is again critically important. Bleaching of the reporters impairs the reliability of the imaging results. During long-term continuous imaging, laser power should be as small as possible, because NSCs are delicate and very sensitive to exogenous perturbations, including photodamage by exposure to a strong laser.

# Oscillatory Expression of bHLH Transcription Factors in Multipotent NSCs

NS cell cultures established from Venus-Hes1 fusion knockin, Venus-Ascl1 fusion BAC Tg, and mCherry-Olig2 fusion BAC Tg mouse strains were used for real-time imaging with confocal fluorescence microscopy. This analysis revealed that the fluorescent fusion reporter proteins with Hes1, Ascl1, or Olig2 exhibited dynamic expression changes in NS cells (**Supplementary Movies 1**–**3**), as observed in the bioluminescence imaging experiments with luciferase-bHLH fusion reporters (Imayoshi et al., 2013b). By quantitative analysis of single cells, the levels of Hes1 and Ascl1 proteins seem to oscillate with a 2–3-h period (**Figures 2A,B**), while Olig2 oscillates with a longer period, such as 5–8 h (**Figure 2C**) (Imayoshi et al., 2013b). Although fluorescent reporter-bHLH proteins showed dynamic expression changes, oscillatory expression was observed more clearly in the bioluminescence imaging experiments with luciferase-bHLH fusion reporters.

Especially, the peak amplitude during signal changes was larger in the case of luciferase-bHLH fusion reporters (Imayoshi et al., 2013b), which is partly because the protein maturation time of luciferase is more rapid than that of fluorescent proteins, such as Venus and mCherry. Our observations revealed that, in actively dividing and multipotent NSCs, bHLH transcription factors are expressed in an oscillatory manner.

The oscillatory expression of bHLH transcription factors raised the possibility that NSCs may change their fate preferences over time. Indeed, when cultured NSCs are sorted by their expression levels of each bHLH transcription factor, their differentiation preferences are correlated with bHLH expression levels (Imayoshi et al., 2013b). For instance, Hes1-high NSCs preferentially differentiate into astrocytes, whereas Ascl1-high and Olig2-high NSCs preferentially differentiate into neurons and oligodendrocytes, respectively. These results suggest that different expression levels of bHLH transcription factors bias the fate choices of NSCs. However, such transient high expression of bHLH transcription factors may not be sufficient for cell fate determination, because these NSCs are still multipotent. Thus, we analyzed how the expression of these bHLH transcription factors changes during cell fate choice.

# Expression Dynamics of bHLH Transcription Factors in Multipotency and Fate Choice

In contrast to oscillatory expression of multiple bHLH transcription factors in self-renewing NSCs, one of the bHLH transcription factors is expressed in a sustained manner during cell fate choice, while the others are repressed (**Figure 3**) (Imayoshi et al., 2013b). For instance, we found that the transient down-regulation of Hes1 expression and the concomitant up-regulation of Ascl1 before cell division bias NSCs toward a neuronal fate choice, and the sustained expression of Ascl1 after cell division irreversibly determines neuronal fate (**Figure 3**). During astrocyte and oligodendrocyte differentiation, the expression of Hes1 and Olig2 is upregulated, respectively, although they still oscillate. However, even during the trough phases, both Hes1 and Olig2 levels are

higher than they are in NSCs, indicating that Hes1 and Olig2 expression continues in a sustained manner during astrocyte and oligodendrocyte differentiation (**Figure 3**). When Hes1 or Olig2 becomes dominant, the expression of the other two factors is downregulated. These results indicate that Hes1, Ascl1, and Olig2 are expressed in an oscillatory manner in multipotent NSCs, and that one of them becomes dominant during cell fate choice (Imayoshi et al., 2013b).

From these analyses, we propose the following model: the oscillatory expression of multiple bHLH transcription factors is correlated with the multipotent and self-renewable state of NSCs, whereas the sustained expression of a selected bHLH transcription factor regulates fate determination (**Figure 3**) (Imayoshi et al., 2013b; Imayoshi and Kageyama, 2014a,b). To validate this model, we applied a new optogenetic method (photo-activatable Gal4/UAS system) to manipulate artificially the expression patterns of bHLH transcription factors using blue light illumination, showing that oscillatory expression activates the proliferation of NSCs, whereas sustained expression induces Imayoshi et al. Oscillatory control of bHLH factors

cell fate determination. Detailed results are reviewed elsewhere (Imayoshi et al., 2013b; Imayoshi and Kageyama, 2014a,b).

# Conclusion

Previous reports have shown some evidence that the bHLH transcription factors Hes1, Ascl1, and Olig2 have multiple functions, such as NSC proliferation, self-renewal, and differentiation. Of course, protein modifications and/or partner co-factor variation may be involved in these different activities of bHLH transcription factors in a complex manner, but our studies revealed that oscillatory versus sustained expression dynamics also contribute to these different functions. More generally, our studies indicated that expression patterns, rather than simply the expression levels, of various transcription factors regulate whether stem cells proliferate or differentiate. In addition to Hes1, Ascl1, and Olig2, other bHLH transcription factors, such as Hes5 and Neurog2, are expressed in an oscillatory manner in NSCs or neural precursor cells (Shimojo et al., 2008; Imayoshi et al., 2013b). Dynamic expression is not necessarily confined to bHLH-type transcription factors, and many kinds of transcription factors are known to be involved in NSC regulation: therefore, it is expected that future imaging and manipulation studies will unveil the unique expression dynamics and their significance of various transcription factors in NSCs.

bHLH transcription factors regulate other kinds of somatic stem cells and pluripotent stem cells. For example, Ascl1 is essential for the differentiation of neuroendocrine cells from epithelial progenitors of the developing glandular stomach (Kokubu et al., 2008), and cyclical Hes1 expression in embryonic stem cells is important for their diverse differentiation abilities (Kobayashi et al., 2009). Therefore, the dynamic control of bHLH transcription factors, including their oscillatory expression, may regulate many regulatory processes of other kinds of stem cells.

Our studies suggest the importance and future possibilities of imaging techniques for the analysis of the expression dynamics of endogenous proteins at the single-cell level by coupling with various cellular events, such as cell cycle progression and cell differentiation (Miyawaki, 2011; Isomura and Kageyama, 2014).

# References

Under the current situation in developmental biology in which many biological processes start to be understood from the viewpoint of systems biology or single-cell biology (Levine et al., 2013; Sanchez and Golding, 2013; Zenobi, 2013), it is expected to become even more important to image subcellular localization and expression dynamics over time at a high resolution while keeping the original expression levels of target proteins.

# Acknowledgments

This work was supported by research grants from the Ministry of Education, Culture, Sports, Science and Technology of Japan (MEXT) and Japan Science and Technology Agency (JST): Grant-in-Aid for Scientific Research on Young Scientists (A) (JSPS 24680035; 15H05570) (II), Challenging Exploratory Research (JSPS 26640011) (II), Innovative Areas (JSPS 26117511; 15H01489) (II), and the JST PRESTO program (II). We also thank the supports by the Human Frontier Science Program, the Nakajima Foundation, the Suzuken Memorial Foundation, the Mochida Memorial Foundation, the Sumitomo Foundation, the Senri Life Science Foundation, the Uehara Memorial Foundation, The Waksman Foundation of Japan INC, the Research Foundation for Pharmaceutical Sciences, the Cell Science Research Foundation, the Brain Science Research Foundation, and Platform for Dynamic Approaches to Living System from the MEXT.

# Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fncel. 2015.00288

Supplementary Movie 1 | Venus-Hes1 expression in self-renewing NSCs. Time in h and min is indicated. Related to Figure 2A.

Supplementary Movie 2 | Venus-Ascl1 expression in self-renewing NSCs. Time in hr min, and sec is indicated. Related to Figure 2B.

Supplementary Movie 3 | mCherry-Olig2 expression in self-renewing NSCs. Time in hr min, and sec is indicated. Related to Figure 2C.

**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 Imayoshi, Ishidate and Kageyama. 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.

# Lipidome of midbody released from neural stem and progenitor cells during mammalian cortical neurogenesis

Yoko Arai † , Julio L. Sampaio, Michaela Wilsch-Bräuninger, Andreas W. Ettinger † , Christiane Haffner and Wieland B. Huttner\*

Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany

#### Edited by:

Chao Deng, University of Wollongong, Australia

#### Reviewed by:

Xiao-Feng Zhao, University of Michigan, USA Yechiel Elkabetz, Tel Aviv University, Israel

#### \*Correspondence:

Wieland B. Huttner, Max Planck Institute of Molecular Cell Biology and Genetics, Pfotenhauerstrasse 108, D-01307 Dresden, Germany<EMAIL_ADDRESS>
#### †Present Address:

Yoko Arai, Institut Jacques Monod, Centre National de la Recherche Scientifique UMR 7592, Université Paris Diderot, Sorbonne Paris Cité, Paris, France; Andreas W. Ettinger, University of California, San Francisco, San Francisco, CA, USA

> Received: 15 June 2015 Accepted: 06 August 2015 Published: 28 August 2015

#### Citation:

Arai Y, Sampaio JL, Wilsch-Bräuninger M, Ettinger AW, Haffner C and Huttner WB (2015) Lipidome of midbody released from neural stem and progenitor cells during mammalian cortical neurogenesis. Front. Cell. Neurosci. 9:325. doi: 10.3389/fncel.2015.00325 Midbody release from proliferative neural progenitor cells is tightly associated with the neuronal commitment of neural progenitor cells during the progression of neurogenesis in the mammalian cerebral cortex. While the central portion of the midbody, a cytoplasmic bridge between nascent daughter cells, is engulfed by one of the daughter cell by most cells in vitro, it is shown to be released into the extracellular cerebrospinal fluid (CF) in vivo in mouse embryos. Several proteins have been involved in midbody release; however, few studies have addressed the participation of the plasma membrane's lipids in this process. Here, we show by Shotgun Lipidomic analysis that phosphatydylserine (PS), among other lipids, is enriched in the released midbodies compared to lipoparticles and cellular membranes, both collected from the CF of the developing mouse embryos. Moreover, the developing mouse embryo neural progenitor cells released two distinct types of midbodies carrying either internalized PS or externalized PS on their membrane. This strongly suggests that phagocytosis and an alternative fate of released midbodies exists. HeLa cells, which are known to mainly engulf the midbody show almost no PS exposure, if any, on the outer leaflet of the midbody membrane. These results point toward that PS exposure might be involved in the selection of recipients of released midbodies, either to be engulfed by daughter cells or phagocytosed by non-daughter cells or another cell type in the developing cerebral cortex.

Keywords: neural stem cells, midbody, shotgun lipidomic analysis, phosphatydylserine, mammalian cortical neurogenesis

# Introduction

At the end of mitosis, after the ingression of the cleavage furrow, a dividing cell is partitioned into two daughter cells that remain connected through a cytoplasmic bridge, the midbody, which is composed of the remnants of the central spindle including a set of microtubule interacting proteins (Glotzer, 2009; Chen et al., 2013). Physical separation of daughter cells is achieved by abscission, which completes cell division/cytokinesis by severing the midbody-bridge, bypassing the central portion of the midbody that contains a dense matrix of antiparallel microtubules and electrondense material (Buck and Tidsale, 1962). Accumulating in vitro evidence from induced pluripotent stem and cancer-derived immortalized cells indicates that the post-abscission midbody, with its characteristic matrix composition, is subsequently either retracted asymmetrically by a daughter cell (single abscission) (Kuo et al., 2011) or released into the extracellular space (double abscission). Alternatively, released midbodies could get attached to the surface of a daughter cell and consequently be engulfed (Ettinger et al., 2011; Crowell et al., 2014). In culture cells, neural stem cells, embryonic stem cells, or cancer-derived cells, which are still responsive to differentiation agents, showed midbody-release into the extracellular space (Ettinger et al., 2011). Potentially, these midbodies might be taken up by non-daughter cells, which could result in a long-range dispersion of midbodies. A similar phenomenon is observed in neuroepithelial cells during mouse cortical development (Marzesco et al., 2005; Dubreuil et al., 2007).

At the onset of neurogenesis, neuroepithelial cells divide asymmetrically and preferentially release their midbody (NE midbody in short) into the extracellular ventricular fluid (Marzesco et al., 2005). The NE midbody release showed a strong correlation with the increase of neurogenesis (Dubreuil et al., 2007). Therefore, an efficient release of midbodies from neuroepithelial cells was postulated as a mechanism to reduce proliferative capacity of neuroepithelial cells. Midbody release ultimately results in loss of both the cytoplasmic and membraneous components present in the midbody including lipids from neuroepithelial cells. While the mechanism of the midbody release from neuroepithelial cells had been addressed through protein functions (Dubreuil et al., 2007), it is still unclear what is the fate of the NE midbody during neurogenesis.

In this study, we characterized the NE midbody for the first time through lipidome analysis, where we observed enrichments of specific classes of phosphatidylserine (PS), phosphatidylethanolamine (PE), and specific species of ceramide (Cer), and triacylglycerols (TAGs). Among these, PS was strikingly enriched in NE midbody. It has been proposed that midbodies with PS, particularly carrying externalized PS on their membrane bilayer, are cleaned up by phagocytes in Caenorhabditis elegans (Chai et al., 2012). This observation suggests that the PS status on the membrane bilayer is associated with the fate of the NE midbody, which may allow deducing its fate. Our data regarding PS status suggests at least two different categories of recipients, either a daughter cell for an engulfment pathway or non-daughter cells for phagocytosis. The distinct subtype of NE midbodies released upon differentiation (Dubreuil et al., 2007; Ettinger et al., 2011) and carry a stem cell marker (Marzesco et al., 2005; Corbeil et al., 2010) may be a novel cue in and contribute to the complexity of the proliferative territories in the developing cerebral cortex.

# Results

#### Midbody Enrichment for Lipidomic Characterization

To enrich midbodies released from neural progenitor cells of the developing mouse embryo ("NE midbody" in short), we collected the ventricular fluid [embryonic cerebrospinal fluid (CF)] of mouse embryos at E11.5 (the number of midbodies is high at this stage) (Marzesco et al., 2005) (**Figure 1A**). In order to separate the midbody particles (∼500 nm) from the

biggest lipid "contaminant," the lipoprotein particles (∼50 nm), which are both present in the ventricular fluid (Marzesco et al., 2005), we subjected the sample to a velocity sucrose gradient. We analyzed all fractions from the gradient by immunoblotting for marker proteins prior lipid analysis in order to check for the quality of the separation (**Figure 1B**). We used the midbody markers: citron rho-interacting kinase (CRIK) (Madaule et al., 1998), prominin-1 and α-tubulin (Dubreuil et al., 2007) in order to identify the fraction enriched in midbody particles in the sucrose fractions. Lipoparticles enriched fractions were identified with the marker apolipoprotein A1 (Furbee et al., 2002) (**Figure 1B**).

Total lipid quantified from cerebrospinal fluid of E11.5 mouse embryos present in the individual fractions 1–9 after velocity sucrose density gradient.

Comparing all fractions in terms of abundance of all markers, we were able to fractionate midbodies (L6) from lipoprotein particles (L1) and cell membranes (L8) (**Figure 1B**). We identified fraction 6 (L6) as the midbody particleenriched fraction due to the relative enrichment of midbody markers CRIK, prominin-1 and α-tubulin and the absence of apolipoprotein A1. While fraction 7 (L7) was also enriched with midbody markers, we did not present this fraction as a midbody fraction because of the increased contamination by apolipoprotein A1 as compared to fraction L6, and considered it being a mixed population of L6 and 8. Fraction 1 (L1) was a relatively pure lipoprotein particle-containing fraction judging from the enrichment of α-tubulin and apolipoprotein A1 and depletion of CRIK and prominin-1. Fraction 8 (L8) was used as a reference of a mixed fraction that included low amounts of midbody and plasma membrane of neural progenitor cells due to the presence of all protein markers (**Figure 1B**). Moreover, we could observe that lipoproteins were by far the most predominant lipid-containing entities present in the ventricular fluid where its purified fraction (L1) outweighs the midbody fraction (L6) by more than seven-fold (**Figure 1C**).

# Midbodies Have a Distinct Lipid Class Composition from Other Cellular Membranes

The lipid analysis of the all fractions confirmed that L1 is indeed enriched in lipoprotein particles. Lipoproteins possess a hydrophobic core composed by neutral lipids such as triacylglycerols (TAG) and cholesterol ester (CE) (Willnow et al., 2007) and we observe that this fraction is the most enriched in these lipid classes (**Figure 2A**). We then see a continuous decrease in the total lipid amount in lower gradient fractions that is accompanied by changes in several lipid classes (**Figures 1C**, **2A**). As expected from fractions more depleted from lipoprotein particles, CEs and TAGs decrease in lower gradient fractions as well as sphingomyelin (SM) (**Figure 2A**). Most importantly, we observe a sharp increase in phosphatidylethanolamine (PE) and phosphatidylserine (PS) in lower gradient fractions that peak precisely at fraction L6 where we observe the purest midbody fraction (L6) followed by a sharp decrease in the following fractions (**Figure 2A**). It is important to note that in the fractions below L6, we also observe a dramatic decrease in TAGs and PE plasmalogens (PE O-) and a slight increase in PC (phosphatidylcholine) suggesting high specificity in terms of lipid class composition of the midbody containing fraction compared to the remaining cell membrane fragments. Importantly, the mean PS amount in L6 was 27-fold higher than in L1 (L6, 2.64 ± 1.26; L1, 0.098 ± 0.11, **Figure 2B**) and 2.4-fold when L6 was compared to the intermediate fraction L8 (L8, 1.11 ± 0.8, **Figure 2B**). Respective to the amount of PE, the mean PE amount in L6 was 9.3 and four times higher than in L1 and L8, respectively (L6, 2.05 ± 0.66; L1, 0.22 ± 0.13; L8, 0.51 ± 0.15, **Figure 2B**).

In summary, we observe high specificity of lipid class composition in the midbody-enriched fraction most notably, PS and PE are highly enriched in the midbody-containing fraction compared to that of lipoprotein and cell membrane mixed fractions.

FIGURE 2 | The Lipidome of NE midbodies. (A) Lipid class profile of all sucrose fraction. The content of individual lipid classes was determined by summing up absolute abundances of all identified species and is expressed as mol %. We consider 100% to be all the membrane lipids, meaning that CE and TAG are excluded from the normalization. CE and TAG amounts are also normalized to the total membrane lipids. P-values were estimated by One-Way ANOVA; \*\*P < 0.01, \*\*\*\*P < 0.0001. Error bars correspond to SD (n = 3). CE amounts are divided by 10 and PC O-, PE, PE O-, PS, and Cer are multiplied by 10 for visualization purposes. CE, cholesterol ester; TAG, triacylglycerol; PC, phosphatidylcholine; SM, sphingomyelin; PI, phosphatidylinositol; Chol, Cholesterol; PC O- and PE O-, plasmalogens; PE, phosphatidylethanolamine; PS, phosphatidylserine, Cer, ceramide. (B) Enrichment of lipid constituency in layer 6 (L6) compared to layer 1 (L1) (L6/L1) and layer 8 (L8) (L6/L8), respectively. The y-axis is on a log2 scale.

# Specific Lipid Species are Enriched in Midbodies Compared to Other Cellular Membranes

It has been previously reported that midbodies collected from HeLa cells ("HeLa midbody" in short) showed the specific accumulation of lipid species of C22–24 ceramides, TAG [16:1, 12:0, 18:1] and PS [18:0–20:4] when compared to cells in cytokinesis (Atilla-Gokcumen et al., 2014). In agreement with the result of the HeLa midbody lipidome, the NE midbody lipidome displays an enrichment of TAG [46:2] in the midbody fraction and a specific enrichment of PS [38:4] and ceramides but in our case, we observed instead a specific increase in C16– 18 ceramides, Cer [36:1:2] (**Figure 3A**). Importantly, we could observe many more species enriched in the midbody fraction. Not only we found TAG [46:2] enriched in the midbody fraction, but also many other TAGs with similar short chain fatty acid compositions (C12–18) in different putative combinations, TAG [46:0], TAG [46:1], TAG [46:2], and TAG [50:4] (**Figure 3A**). We could also observe that most lipid species belonging to PS and PE irrespective of the fatty acid saturation and chain length level display a strong enrichment in the midbody fraction (**Figures 3B,C**). Interestingly, specific PE O- species, namely with longer and more unsaturated fatty acids, PE O- [38:6] and PE O- [40:6] are enriched in the midbody fraction (**Figure 3C**). Although PC as a class is not changing significantly between the different fractions, we can observe some changes in lipid

species. Shorter and more saturated PC lipid species (C30–34) are enriched in midbodies, PC [30:0], PC [32:0], PC [32:1], and PC [34:1], while longer and more unsaturated lipid species are depleted in the midbody fraction, PC [36:2], PC [36:4], and PC [38:4] (**Figure 3D**).

In conclusion, we could reproduce the previous published results on lipids enriched in the midbody of Hela cells but identified new midbobody specific lipids, namely within Cer, PS and TAG lipid classes, PE as a lipid class and long and unsaturated PE O- and a remodeling of the PC lipid class to shorter and more saturated lipid species in the midbody.

# NE Midbodies Carry Internalized and Externalized PS

From the lipidome analysis, we observed that mainly PC is the most depleted membrane lipid classes (aprox. 8 mol %) while PS and PE are the most enriched lipid classes in NE midbody (almost 5 mol % in total) compared to cell membranes (L8) (**Figures 3B–D**). If we take into consideration the plasma membrane lipid asymmetry where PC is on the extracellular leaflet of the plasma membrane at the cell surface, and both PS and PE face the cytoplasmic leaflet (Verkleij et al., 1973), it raises the interesting possibility that PS and/or PE might be externalized to keep the membrane integrity. In C. elegans, externalization of PS on midbodies released from Q neuroblasts was shown to be an engulfment signal for the midbody via clearance by phagocytes (Chai et al., 2012). These lines of evidence suggest that the status of PS externalization on the midbody might be a mark to infer the fate and/or recipient cell of released midbodies.

Therefore, we assessed the leaflet localization of PS in midbody membranes, taking advantage of its specific interacting partner, Annexin V (Schutte et al., 1998). To this end, we injected Annexin V directly coupled with Cy5 (Annexin V-Cy5) into the cerebral fluid of mouse embryos at the level of the forming cortex (E11.5), allowing binding for 30 min, followed by fixation and analysis by immunofluorescence. At this early stage of neurogenesis, the fluid filling the ventricular space contains a high amount of released NE midbodies. In this experimental setup, binding of Annexin V-Cy5 to particles is expected only if PS was externalized on the outer leaflet of the membrane bilayer. We confirmed Annexin V-Cy5 positive particles in the ventricular fluid as NE midbodies by a counterstaining with prominin-1, a midbody marker (Marzesco et al., 2005) (**Figures 4A–D**). In addition to particles in the lumen, we also detected Annexin V-Cy5 staining at the apical plasma membrane of neuroepithelial cells, which is the site of cytokinesis midbody formation between nascent daughter cells (**Figure 4C**). The presence of Annexin V-Cy5 and prominin-1 double positive midbody structures on the apical surface was confirmed by immunogold labeling and electron microscopy (**Figures 4E–J**). We found comparably electron-dense, ∼500 nm–large structures labeled individually or together by antibodies against prominin-1 or Cy5 (**Figures 4E–J**). These results indicate that PS externalization could occur at the end of cytokinesis of neuronal progenitor cells.

NE midbodies show three distinct populations defined by their decoration with Annexin V-Cy5 and presence of prominin-1. At the apical surface of the ventricular zone, we observed prominin-1 positive and Annexin V-Cy5 negative particles as a major population (57%, **Supplementary Figure 1**) and prominin-1 and Annexin V-Cy5 double positive particles as a second population (41%, **Supplementary Figure 1**). A third minor population (2%) contained only Annexin V-Cy5. Interestingly, in the CF, prominin-1, and Annexin V-Cy5 double positive particles became the major population (55%, **Figure 4K**)

and the prominin-1 positive and Annexin V-Cy5 negative particles were the second population (40%, **Figure 4K**). The third minor population (5%) contained only Annexin V-Cy5 was still observed. Taken together, these results strongly suggest that a fraction of neural progenitor cell midbodies externalized PS prior to their release in the ventricular fluid and ultimately both PS-externalized and -internalized midbodies were found after cytokinesis.

# PS Flipping Occurs Upon the Detachment of the Midbody after Cytokinesis

In order to determine the dynamics of flipping of PS on the membrane, we analyzed dividing neuroblastoma cells (Neuro-2a), which have been shown to have the capacity of both releasing and engulfing midbodies (Ettinger et al., 2011). We performed live-cell time-lapse microscopy using a transgenic Neuro-2a cell line expressing GFP- tagged mitotic kinesin-like protein-1 (MKLP1) (Poser et al., 2008; Ettinger et al., 2011), a core midbody and central spindle protein (Hu et al., 2012). This cell line allows the visualization of the forming and released midbody (Neuro2a midbody, in short). To visualize externalized PS, we added Annexin V-Cy5 to the culture medium at the beginning of the movie. We assessed PS externalization by appearance of the Cy5 signal upon binding of Annexin V-Cy5. Midbodies were identified as bright spots in the GFP channel.

As reported previously (Ettinger et al., 2011), MKLP1- GFP localized to the central portion of the midbody in Neuro-2a cells. PS was not externalized during the telophase (**Figure 5A**, "T"). The midbody was subsequently detached from the central portion of the midbody bridge between daughter cells after either a first or second abscission event of the cytoplasmic membrane. This detachment was identified by the displacement of the midbody from the point of contact of the daughter cell to one of the two daughter cells and PS was externalized at this moment, as determined from appearance of a Cy5 spot (**Figure 5A**, "A"). Approximately 1 h after the detachment, the midbody was released from a daughter cell (**Figure 5A**, "R"). These results indicate that PS was flipped out and exposed to the extracellular space at the end of the cytokinesis before midbody release (see Supplementary Movie 1).

We quantified Annexin V and MKLP1-GFP double positive midbodies in Neuro-2a MKLP1-GFP cell cultures in order to understand how widespread this phenomenon is (**Figures 5B–E**). During late telophase, a midbody bridge is formed between the nascent daughter cells. MKLP1-GFP was recruited to the

as z-stacks every 0.38-µm and are maximum intensity projections. Scale bars, 10µm. (B) Examples of bilateral midbodies. Fluorescence

central portion of the midbody (**Figure 5B**) as confirmed by immunostaining for α-tubulin to highlight the central spindle microtubules and for Aurora B kinase which localizes to the midzone in anaphase and decorates the inner part of the central spindle in telophase (**Supplementary Figure 2A**) (Hu et al., 2012). We almost never detected Annexin V-Cy5 staining in these newly formed midbodies (about 3.5% of total bilateral midbodies, **Figures 5B,E**). Instead, Annexin V was present in midbodies either adhering to one of the daughter cells after a first abscission or on interphase cells (**Figures 5B,C**, white arrowheads and see **Supplementary Figure 2B**). The majority of such monolateral midbodies were Annexin V-Cy5 positive (≈66% of total monolateral midbodies, **Figure 5E**). Fully released Neuro2a midbodies contained Annexin V-Cy5 in the majority of cases (≈90% of total released GFP positive particles, **Figures 5D,E**). Taken together, PS exposure occurred before the complete release of the midbody and it likely occurred after the first abscission step.

#### PS Status on HeLa Midbodies

and 137 for released); error bars indicate SD.

A previous study indicated that half of the total Neuro-2a midbodies are retained on daughter cells and the rest are released into the extracellular space (Ettinger et al., 2011). Conversely, in HeLa cells, midbodies were described to be engulfed by a daughter cell (Crowell et al., 2014). Therefore, we asked whether HeLa midbodies carry externalized PS. Similarly to the analysis of Neuro2a cells, we used a transgenic HeLa cell line expressing MKLP1-GFP to follow the midbody and determined the presence of PS in the outer leaflet of the midbody membrane with Annexin V-Cy5 (**Figure 6**). Midbody bridges formed during telophase were identified by recruitment of MKLP1-GFP and confirmed by immunostaining for α-tubulin and Aurora B (**Supplementary Figure 3**). In both monolateral inherited midbodies (**Figure 6A**) and in newly formed/bilateral midbodies (**Figure 6B**), we hardly detected PS externalization. These results suggest that PS exposure to the extracellular environment is a characteristic feature of neural-related cell lines and neuronal progenitor cells.

(total number of counted particles, 243 for bilateral, 234 for monolateral

# Discussion

The present study is the first quantitative analysis of the lipid composition of released NE midbodies from mouse embryonic CF. A comparison of lipidome data from HeLa and NE midbodies showed a common enrichment of several lipid species (Atilla-Gokcumen et al., 2014). Among them, the enrichment in specific PS, TAG and Cer species was observed both in NE and HeLa midbodies compared to both lipoprotein particles and dividing cells. In comparison to midbodies from HeLa cells, NE midbodies displayed a wider extent of enrichment of other lipid classes and specific lipid species. This apparent divergence may reflect the functional complexity of the NE midbody in developing mouse embryos, in contrast to cancerderived, cultured cells like HeLa. It has been shown previously that cultured cells showed a re-arrangement of protein levels involved in metabolic pathways at the expense of other pathways as compared to primary cells, which seems to be required for the adaptation to the cultured conditions (Pan et al., 2009). Particularly relevant for this study is the down-regulation of proteins involved in fatty acid metabolism in cell lines (Pan et al., 2009), a fact that can explain why we identified more varying lipid species in NE midbody when compared to HeLa midbody.

What could be the relevance of the combined enrichment of all the reported lipids for the midbody structure and function? Interestingly, both Ceramide species (C16 and C24-ceramide) identified in our and the previous study (Atilla-Gokcumen et al., 2014), respectively have been shown to contribute for apoptosis (Seumois et al., 2007). Moreover, the enrichment and exposure of PS to the outer leaflet of plasma membrane, a feature observed in midbody membranes, is a common feature of many apoptotic cells (Fadok et al., 1998). These two observations put together suggest that released midbody membranes might contain a potent apoptotic signal and, upon its releasing from the neural stem and progenitor cells and its engulfment by neighboring cells, it might alter the cell fate of both. The strong enrichment in unsaturated PE species in the midbody, is in line with its recognized fusogenic properties and the finding that PE species are exposed on the cell surface specifically at the cleavage furrow during late telophase of cytokinesis (Emoto et al., 1996). PE, due to its unique geometry (inverted cone) which is enhanced by the increase of the degree of unsaturation, in that it forms a non-bilayer structure, is thought to promote rapid phospholipid trans-bilayer movement (Ellens et al., 1989) and stalk membrane formation (Markin et al., 1984) facilitating membrane fusion and fission processes in cellular membranes a process required for midbody release after cytokinesis.

Cells in cytokinesis have been shown to possess more resilient cell membranes requiring three-fold higher forces to provoke their disruption when compared to cells in interphase (Atilla-Gokcumen et al., 2014). This observation is in line with the increase in relative saturation of the most abundant phospholipid in mammalian cells, PC (**Figure 2D**). We observe enrichment in PC species containing 0 and 1 unsaturations and a decrease in PC species containing two or more unsaturations (**Figure 2D**). Unsaturation level of lipids is known to alter dramatically the physical properties of cell membranes by changing the membrane fluidity and order [Gennis RB, Biomembranes: Molecular structure and function (Springer Advanced Texts in Chemistry), 1989]. The specific enrichment in short chain TAGs is intriguing. TAGs in general are known to be metabolic energetic reservoirs in cells but the putative different functions of TAGs with specific fatty acid composition are still unknown.

Together, the lipid specificity observed in midbodies, seems to be intimately related to its structure and function. From it, we can conclude that midbodies possess relatively sturdy membranes due to the decrease of saturation of PC species, they are enriched in fusogenic lipids, unsaturated PEs, which facilitate fission and release of midbody, and are also enriched in potent signaling lipids, specific Ceramides and PS species that might influence neighboring cells if engulfed by them. All these observations put together highlight the importance of lipid compositional complexity, specificity and cellular localization in cell biology and how it can be directly associated with structural and functional aspects of cellular processes (Simons and Sampaio, 2011).

Although differences in lipid composition exist, common lipid species suggest shared characteristics between NE and HeLa midbodies. In HeLa cells, midbodies are known to be engulfed by a daughter cell through an actin-dependent phagocytosislike mechanism (Crowell et al., 2014) while the release of midbody from neural-derived cells (Ettinger et al., 2011) seems not compatible with the engulfment by one of the daughter cells. PS is a structural phospholipid, which exists in the cytoplasmic leaflet of the plasma membrane bilayer and whose externalization has been associated with phagocytosis during apoptotic body clearance (Fadok et al., 1998), retinal rod and cone photoreceptors clearance (Kevany and Palczewski, 2010) and also in midbody clearance in C. elegans (Chai et al., 2012). In above examples, the phagocytosis of PS-externalized midbodies was carried out by non-daughter cells. Hence, it is possible that PS-externalized NE midbodies may be engulfed by phagocytes in the developing mouse embryo. It has been proposed, however, that other cell types in the developing mouse central nervous system such as microglia, astrocytes and even neural progenitor cells or neurons have the potential to behave as phagocytes due to the expression of phagocytic receptors (Sokolowski and Mandell, 2011). One of the phagocytic receptors, Gas6, which binds to PS, is indeed expressed highly in proliferative neural progenitor cells compared to neurogenic progenitor cells (Arai and Huttner, unpublished data). These lines of evidence support the idea that PS-externalized NE midbodies could be phagocytosed by sister neural progenitor cells through a phagocytosis-related mechanism using Gasp6 signaling. We previously reported that midbodies carrying the stem cell marker prominin-1 (Marzesco et al., 2005) were released from proliferative neural progenitor cells, which may contribute to their loss of stemness (Dubreuil et al., 2007). One possibility is that PS-externalized midbody could be engulfed by proliferative neural progenitor cells to maintain a balance for the stem cell capacity or serving as a means of communicating between the proliferative territories of the developing mouse brain. In addition to the PS externalized NE midbody, we also observed PS internalized NE midbodies (**Figure 4K**), which may reflect the heterogeneity of neural progenitor cells providing these two different types of midbodies.  It would be interesting to identify which type of neural progenitor cells preferentially release either PS-externalized or -internalyzed midbodies.

What could be the fate of the biological components of NE midbodies after cells take them up? The engulfed midbody should be digested in lysosomes into monomolecular species, which are recycled if necessary. Fatty acids from the PS species of the NE midbody could also be recycled, if any, it participates on any metabolic pathways. Amongst the PS species identified in our lipidomics, the presence of omega-3 and 6 fatty acids that cannot be produced de novo have been strongly suggested, for instance, (PS [38:4] = PS [18:0–20:4], arachidonic acid [20:4]) (Sampaio et al., 2011). Therefore, the recycling of midbodies could participate to the essential lipid metabolism during cortical development, participating to the maintenance of a lipid species pool for controlling proper neurogenesis.

# Methods

## Animals

All mouse embryos used were wildtype C57BL/six mice obtained by overnight mating of wildtype C57BL/six males and females. Noon of the day on which the vaginal plug was observed was defined as embryonic day (E) 0.5. All animal studies were conducted in accordance with German animal welfare legislation, and the necessary licenses obtained from the regional Ethical Commission for Animal Experimentation of Dresden, Germany.

# Enrichment of the Midbody Fraction Using Velocity Sucrose Density Gradient Centrifugation

E11.5 C57BL/6 mouse embryos were dissected in ice-cold 150 mM NH4Ac dissolved in lipid mass spectrometry-grade water (Merck; LiChrosolv grade). CF was collected as described (Marzesco et al., 2005). 1–2µl of CF per embryo were collected and pooled in a 1.5 ml Eppendorf tube (Eppendorf) on ice. Seventy-six micro liter of CF were mixed with 4µl of 100x proteinase inhibitor mix (Roche) and 320µl of 2.5 M sucrose in water to yield a total of 400µl of CF mixture at a final concentration of 2 M sucrose. The CF mix was transferred into the bottom of an ultraclean ultracentrifugation tube (Beckman coulter). The gradient was prepared by adding layers of less dense sucrose solutions (400µl of 1.6 M, 400µl of 1 M, 300µl of 0.3 M, and 300µl of 0.1 M) upon one another. The samples were centrifuged in a TLS55 swinging rotor (Beckman coulter) at 50,000 rpm for 18 h at 4◦C with a no deceleration profile. Immediately after the centrifugation, 180µl fractions were collected into 1.5 ml Eppendorf tubes on ice. Ten micro liter of each fraction were used to determine the sucrose concentration in the fraction, 85µl were used each for Western blotting and lipid mass spectrometry. In case of electron microscopy (EM) analysis, 10µl of each fraction were fixed overnight at 4◦C in 0.5% of EM grade glutaraldehyde/4% paraformaldehyde in 0.12 M sodium phosphate buffer, pH 7.2.

# Protein and Lipid Precipitations from Sucrose Samples

For immunoblotting, samples were subjected to methanolchloroform precipitation by mixing samples in 1:4:1:3 ratio (sample:methanol:chloroform:water). Protein was precipitated by gently vortexing the tube followed by centrifugation at 13,000 rpm for 5 min at room temperature (r.t.). The upper layer was removed without touching the interface layer. Twohundred and fifty-five liter of methanol were added (3 × volume of the original sample volume, 85µl). Proteins were pelleted by centrifugation at 13,000 rpm for 5 min at r.t. and resuspended in 2x concentrated SDS Laemmli sample buffer. Protein samples were denatured by boiling at 95◦C for 5 min and stored at −20◦C.

For lipid extraction, 85µl sample from each fraction were transferred into a 1.5 ml Eppendorf tube filled with 157 mM of ammonium bicarbonate (NH4HCO3) up to 1.5 ml. Particles were pelleted in the TLA55 fixed-angle rotor (Beckman coulter) at 30,000 rpm for 4 h at 4◦C. Removed NH4HCO3 and pelleted lipid samples were frozen in liquid nitrogen and stored at −80◦C.

#### Lipid Extraction for Lipid Mass Spectrometry

Lipid samples were mixed with 200µl of 157 mM of NH4HCO3 and vortexed at 1400 rpm at 4◦C. Ten micro liter of an internal standard lipid mix (Avanti Polar lipids; PC, PE, PS, PG, PA, Cer, SM, GalCer, LacCer, Chol, and DAG) was added to each sample. To extract all lipids, 1 ml of a 2.75:1 mixture of chloroform/methanol was added to each tube and vortexed at 1400 rpm for 1 h at 4◦C. Fractions with high sucrose content (>1 M) were "pre-washed" with 200µl of MS grade water before the extraction to remove excess sucrose. After extraction, sample tubes were centrifuged at 1000 g for 2 min at 4◦C. Bottom phases were transferred into fresh 1.5 ml Eppendorf tubes (10:1 lipid fraction). The organic solvent was evaporated using a vacuum centrifuge. Dried samples were re-suspended with 100µl of a 1:2 mixture of chloroform/methanol and vortexed for 10 min at 4◦C. This re-suspension solution was mixed with MS compatible polarity specific buffers to optimize for ionization of specific lipids as described previously (Carvalho, 2012). The data was acquired on a LTQ-Orbitrap using high mass resolution Rm − z = 400 = 100, 000 and the spectra was analyzed with LipidXplorer software (Herzog et al., 2011, 2012).

#### Immunofluorescence, Electron Microscopy, Annexin V-Cy5 Injection, and Image Acquisition

Living embryos at E11.5 were dissected out of the uterus with the placenta intact and transferred into 1x Tyrode solution (Sigma-Aldrich). AnnexinV-Cy5 solution (BioVision) was diluted to 1:300 in 1x binding buffer (10 mM Hepes-NaOH pH7.4 containing 140 mM NaCl and 2.5 mM CaCl2) and injected into telencephalic vesicles of living embryos (1x concentrated binding buffer was used as control). Injected embryos were incubated at r.t. for 30 min (room air and protected from light). Whole embryos were fixed for 20–25 h at 4◦C in 4% paraformaldehyde in 0.12 M sodium phosphate buffer, pH 7.2. For cryosectioning, embryos were equilibrated for 12–24 h at 4 ◦C in 30% sucrose in PBS, embedded in Tissue-Tek O.C.T compound (Sakura Finetek), and stored at −20◦C. Cryostat sections (10µm) were rehydrated with PBS, quenched with 50 mM NH4CL for 30 min, washed with 0.01% Digitonin (Sigma-Aldrich) in PBS for 30 min and blocked with 3% BSA in 0.01% Digitonin/PBS for 30 min at r.t. Sections were washed with 1% BSA in 0.01% Digitonin/PBS and incubated overnight at 4 ◦C with first primary antibody, followed by incubation for 1 h at r.t. with fluorescently labeled secondary antibodies and DAPI (4′ ,6-diamidino-2-phenylindole, Roche) in 1% BSA in 0.01% Digitonin/PBS. Sections were mounted in Mowiol. For electron microscopy, E11.5 dorsal telencephalon was analyzed by transmission EM, with pre- or post-embedding prominin-1 immunolabeling.

The following primary antibodies were used: rabbit anti-Prominin-1 [αD, MPI-CBG; 1:20,000 for immunoblotting (IB)]; goat antibody against apolipoprotein A1 (Biodesign; 1:1000); mouse monoclonal antibodies against CRIK [BD bioscience; 1:200 for immunofluorescence (IF) and 1:4000 for IB], α-tubulin (Sigma; 1:300 for IF and 1:60,000 for IB) and Aurora B (BD bioscience; 1:300) and rat anti-Prominin-1 (13A4, MPI-CBG; 1:300 for IF). Secondary antibodies were: horseradish peroxidasecoupled antibodies (Jackson laboratory), Alexa Fluor 488, 555, and 647 labeled IgG antibodies (Invitrogen).

#### Live-cell Time-lapse Microscopy

The protocol for this time-lapse microscopy has been described previously (Ettinger et al., 2011). Briefly, Neuro-2a MKLP1- GFP cells were grown on chambered coverslips (Lab-Tek, Nunc) in DMEM-GlutaMax medium supplemented with 10% FCS and incubated for 24 h at 37◦C with 5% CO2. The medium was replaced with Annexin V-Cy5 containing medium (1:5000) and cells were imaged with a Zeiss Axiovert 200 M inverted epifluorescence microscope equipped with a motorized stage, incubator, humidifier, and carbon dioxide supply (Visitron Systems, Germany). Bright-field and fluorescence images were acquired using a C-Apochromat 40x water immersion objective (NA 1.4) and recorded with a Roper Scientific Cool SNAP ES CCD camera using MetaMorph software. Multiple fields were acquired in parallel per experiment at 12 min intervals for up to 24 h.

#### Annexin V-Cy5 Incubation with the Neuro-2a-MKLP1 Transgenic Cell Line

Neuro-2a MKLP1-GFP cells were grown on poly-L-lysine coated coverslips in DMEM-GlutaMax medium supplemented with 10% FCS and incubated for 24 h at 37◦C with 5% CO2. The medium was replaced with Annexin V-Cy5 containing medium (1:2500), incubated for 10 min at r.t. protected from light, and fixed with 4% paraformaldehyde in 0.12 M sodium phosphate buffer, pH 7.2. Cells were further processed for immunofluorescence like sections of mouse embryos.

# Author Contributions

YA designed and performed the experiments, analyzed the data and co-wrote the manuscript, JS designed and performed lipid mass spectrometry, analyzed the data and co-wrote the manuscript, MB performed EM and co-wrote the manuscript, AE helped and performed time-lapse imaging, and co-wrote the manuscript, CH performed experiments and WH analyzed data, co-wrote the manuscript and supervised project. The authors declare that they have no competing financial interests.

# Acknowledgments

We thank the animal and light microscopy facilities of MPI-CBG for excellent support, Drs. Vitary Matiash, Dominik Schwudke, Andrej Shevchenko, and Kai Simons for helpful discussion at the beginning of the lipid mass spectrometry and project. YA was supported by ARC (Association pour la Recherche sur le Cancer), FRM (Fondation pour la Recherche Médicale), and JSPS (Japan Society for the Promotion of Science) and WH was supported by grants from the DFG (SFB 655, A2; TRR 83, Tp6) by the DFGfunded Center for Regenerative Therapies Dresden, the Fonds der Chemischen Industrie and ERC.

# Supplementary Material

The Supplementary Material for this article can be found online at: http://journal.frontiersin.org/article/10.3389/fncel. 2015.00325

# References

Supplementary Figure 1 | Phosphatidylserine is on the outer-leaflet of the apical plasma membrane of the neuronal progenitor cells. (A,B) Micrographs of mouse embryonic E11.5 coronal cryosections of dorsal telencephalon injected with Annexin V-Cy5 binding buffer. Sections were consecutively stained by immunofluorescence for prominin-1 (green) and counterstained with DAPI (white). Images are z-projections of 32 consecutive 0.38-µm optical sections. (B) Higher magnifications of the apical plasma membrane of the ventricular zone [boxed region indicated in (A)]. Red-white arrows, Annexin V-Cy5 single positive particles, green arrows, prominin-1 single positive midbody particles, yellow arrows, Annexin V-Cy5 and prominin-1 double positive midbody particles. Scale bars, 50µm (A) and 10 µm (B). (C) Percentage of three different types of particles on the apical plasma membrane of the ventricular zone, classified by decoration with prominin-1 positive and Annexin V-Cy5 positive signal. Data are the mean of six 213-µm wide fields from three different embryos and litters; total number of counted particles, 455 for prominin+ AnnexinV–, 354 for prominin+ AnnexinV+ and 16 for prominin– AnnexinV+; error bars indicate SD; ∗P < 0.05, ∗∗∗∗P < 0.0001; unpaired t-test.

Supplementary Figure 2 | Expression of midbody markes in MKLP1-GFP Neuro-2a cells. (A,B) Fluorescence for MKLP1-GFP (green),

immunofluorescence for α-tubulin and Aurora B kinase (red) in Neuro-2a MKLP1-GFP cells during telophase (A) and interphase (B) combined with DAPI staining (white). Images are maximum intensity projections of z-stacks of optical sections taken every 0.38µm. Insets, magnifications of bilateral (A) and monolateral (B) midbodies. Scale bars, 10 µm and 1 µm (inset).

Supplementary Figure 3 | Expression of midbody markers in MKLP1-GFP HeLa cells. (A) Fluorescence for MKLP1-GFP (green) and immunofluorescence for α-tubulin in telophase (2 upper panels) and interphase (lower panels) cells combined with DAPI staining (white). Scale bars, 10µm. (B) Fluorescence for MKLP1-GFP (green) and immunofluorescence for Aurora B kinase in telophase (upper panels) and interphase (lower panels) cells having midbody combined with DAPI staining (white). Scale bars, 10µm. Insets, magnifications of newly formed midbodies with telophase-bridge between the two daughter cells intact [upper panels in (A,B)] or inherited, monolateral midbodies with no apparent cytoplasmic bridge to the other daughter cell [lower panels in (A,B)]. Scale bars, 1 µm.

**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 Arai, Sampaio, Wilsch-Bräuninger, Ettinger, Haffner and Huttner. 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.

# Regulation of cerebral cortex development by Rho GTPases: insights from *in vivo* studies

# *Roberta Azzarelli 1, Thomas Kerloch2,3 and Emilie Pacary2,3\**

*1 Department of Oncology, Hutchison/MRC Research Centre, Cambridge Biomedical Campus, University of Cambridge, Cambridge, UK*

*2 Institut National de la Santé et de la Recherche Médicale U862, Neurocentre Magendie, Bordeaux, France*

*3 Institut National de la Santé et de la Recherche Médicale, Physiopathologie de la Plasticité Neuronale, Université de Bordeaux, Bordeaux, France*

#### *Edited by:*

*Takeshi Kawauchi, Keio University School of Medicine / Precursory Research for Embryonic Science and Technology (PRESTO), Japan Science and Technology Agency (JST), Japan*

#### *Reviewed by:*

*Troy Ghashghaei, North Carolina State University, USA Angeliki Louvi, Yale, USA*

#### *\*Correspondence:*

*Emilie Pacary, Neurogenesis and Physiopathology Group, INSERM U862 - Neurocentre Magendie, 146 rue Léo Saignat, 33077 Bordeaux cedex, France e-mail<EMAIL_ADDRESS>
The cerebral cortex is the site of higher human cognitive and motor functions. Histologically, it is organized into six horizontal layers, each containing unique populations of molecularly and functionally distinct excitatory projection neurons and inhibitory interneurons. The stereotyped cellular distribution of cortical neurons is crucial for the formation of functional neural circuits and it is predominantly established during embryonic development. Cortical neuron development is a multiphasic process characterized by sequential steps of neural progenitor proliferation, cell cycle exit, neuroblast migration and neuronal differentiation. This series of events requires an extensive and dynamic remodeling of the cell cytoskeleton at each step of the process. As major regulators of the cytoskeleton, the family of small Rho GTPases has been shown to play essential functions in cerebral cortex development. Here we review *in vivo* findings that support the contribution of Rho GTPases to cortical projection neuron development and we address their involvement in the etiology of cerebral cortex malformations.

**Keywords: Rho GTPases, cerebral cortex, neuronal development, cortical malformations, GAP, GEF**

#### **INTRODUCTION**

In humans, the cerebral cortex is responsible for the processes of thought, perception and memory and serves as the seat of advanced motor functions, social abilities and language. These functions relies on the proper development of two main populations of neurons, projection or pyramidal neurons, which are glutamatergic and excitatory, and interneurons, which are GABAergic and inhibitory. Neurons in the cerebral cortex are arranged into six distinct layers different in terms of connectivity, gene expression profile and birthdate (Molyneaux et al., 2007). Projection neurons originate from progenitors located in the cortex whereas interneurons are born in the ventral domains of the telencephalon and then migrate tangentially to reach the cortex (Kriegstein and Noctor, 2004).

After closure of the neural tube, the epithelium lining the ventricles becomes a specialized neuroepithelium. It consists of a single sheet of progenitor cells, called neuroepithelial cells (NEs). At early developmental stages, around E10 in mouse, NEs along the dorsal surface of lateral ventricles self-renew to expand the progenitor pool and then convert into cells expressing glial markers such as the astrocyte-specific glutamate transporter (GLAST) and brain lipid-binding protein (BLBP), the radial glial cells (RGs). The asymmetric divisions of these RGs are responsible for producing cortical projection neurons either directly or indirectly through intermediate progenitor cells (IPs) or outer radial glial cells (oRGs). Newborn projections neurons then migrate radially in a step-wise fashion to their final destination using RG fibers as a scaffold and finally undergo terminal differentiation to transmit and receive information. As neurogenesis progresses, diverse subtypes of projection neurons are generated sequentially and their migration occurs in an inside-out manner: neurons generated first occupy the deepest layers of the future six layered neocortex whereas later born neurons by-pass earlier born neurons and settle in more superficial layers (Gupta et al., 2002).

Unlike the projection neurons, the inhibitory interneurons of the cerebral cortex are generated from distinct progenitors in the germinal zones of the ventral telencephalon, mainly within the medial and caudal ganglionic eminences. Interneurons then undertake tangential migration toward the cortex using different routes according to the time and place of birth. Upon arrival in the cortex, interneurons switch their mode of migration from tangential to radial and reach their destination layer based on their molecular subtype, origin and birthdate, on cortical cues and make local connections with pyramidal cells (Bartolini et al., 2013; Guo and Anton, 2014).

The development of each category of cortical neurons is thus a multistep process, which involves dramatic morphological changes at each step of the process. These changes are mediated by an extensive and dynamic remodeling of the cytoskeleton. Among the major regulators of cytoskeletal properties, the small signaling molecules of the Rho GTPase family play essential functions in cerebral cortex development. In this review, we will address the role of Rho GTPases in cortical projection neuron development, focusing on *in vivo* studies, and discuss how dysfunctional Rho GTPase signaling may contribute to cortical malformations.

## **THE Rho GTPase FAMILY**

The Rho family of GTPases represents a subgroup of the Ras superfamily of small GTP binding proteins (Heasman and Ridley, 2008). The most extensively studied members of the Rho family are RhoA (Ras homologous member A), Rac1 (ras related C3 botulinum toxin substrate 1) and Cdc42 (cell division cycle 42) but this family actually includes 20 members which are subdivided into 8 subgroups based on amino-acid sequence similarities (**Figure 1**).

Like other small GTP-binding proteins of the Ras superfamily, most Rho GTPases cycle between GTP (active) and GDP (inactive)—bound states. The GDP/GTP cycle is promoted by the activity of two classes of molecules, guanine nucleotide exchanging factors (GEFs) and GTPase activating proteins (GAPs). GEFs facilitate the exchange of GDP with GTP, resulting in protein activation. GAPs instead stimulate the intrinsic enzymatic activity of the GTPases, which promotes hydrolysis of GTP into GDP. GAP activity therefore ends the cycle and returns the GTPases in their inactive state (Bos et al., 2007) (**Figure 2**). Over 80 GEFs and more than 70 GAPs have been reported, suggesting that Rho GTPase regulation is complex. In addition, Rho GTPases can bind to proteins known as guanine-nucleotide dissociation inhibitors

**FIGURE 1 | Members of the Rho GTPase family.** The column identity indicates the percentage of amino-acid sequence identity of a specific Rho GTPase compared with the first member of the corresponding subfamily (Heasman and Ridley, 2008). Among the atypical members, Rnd1, Rnd2, Rnd3, RhoH, RhoBTB1, and RhoBTB2 lack amino acids in the GTPase domain that are critical for GTPase activity, they are thus constitutively bound to GTP and do not detectably hydrolyse GTP (deficient Rho GTPase domain in dark blue). RhoV and RhoU harbor GTPase activity but they are atypical in this family as they display a high intrinsic guanine nucleotide exchange activity and are predominantly in the GTP-loaded conformation. In the C-terminal domain, the Hyper Variable Region (HVR in red) differs not only between the Rho GTPase subclasses but also within the same subclass in terms of the presence of either a polybasic region or a palmitoylation site (Roberts et al., 2008). The polybasic region and palmitoylation site present in the HVR are involved in targeting the GTPases to plasma membrane or

endomembrane compartment. The C-terminal CAAX-box (C, cysteine; A, Aliphatic Amino acid; X, any amino acid; in orange) contains a cysteine residue, which is crucial for prenylation that adds a farnesyl or geranylgeranyl group, enhancing the interaction with membranes and very often defining the localization to specific membrane compartments. The CAAX with a ∗ indicates that RhoV does not seem to have a functional CAAX box and the CAAX motif of RhoU is apparently not undergoing prenylation (Aspenstrom et al., 2007). The Rif members have a N-terminal extension (in yellow) that is unique to this subgroup. RhoUV proteins display a proline rich motif (PRM, in green), which is also present in RhoBTB1 and RhoBTB2. RhoBTB proteins contain two Broad complex/Tramtrack/Bric-a-brac domains (BTB). RhoBTB family members harbor different domains involved in protein-protein interaction (in gray: Coiled Coil, CC, only present in RhoBTB1 and PEST domain only present in RhoBTB2) and they display a nuclear localization signal (NLS, in pink).

(GDIs). RhoGDIs sequester RhoGTPase in their inactive state and protect them from degradation (Boulter et al., 2010) (**Figure 2**). When bound to GTP, Rho GTPases exhibit the correct structural conformation to interact with effectors and initiate downstream signaling to regulate actin and microtubule components of the cytoskeleton (Jaffe and Hall, 2005) (**Figure 2**). However, some members of the family do not follow this classical scheme of activation and are described as atypical. These atypical Rho GTPases are predominantly GTP bound, owing either to aminoacid substitutions at residues that are crucial for GTPase activity (for example in Rnd proteins) or owing to increased nucleotide exchange (for example in RhoU). Therefore, their expression, localization, stability and phosphorylation control their activity rather than the GDP/GTP switch (Aspenstrom et al., 2007) (**Figure 1**).

Experimental data indicating the importance of the Rho family of small GTPases in cerebral cortex development have been accumulated over the past few years. Much of our understanding on their role comes from *in vitro* studies (Govek et al., 2005). Nevertheless, in the last years, the use of conditional mutant mice and the development of techniques such as *in* *utero* electroporation have allowed to highlight and clarify their functions *in vivo*.

#### **EXPRESSION OF Rho GTPases IN THE DEVELOPING CEREBRAL CORTEX**

Most of these *in vivo* studies have focused on RhoA, Rac1, and Cdc42, and more recently on Rnd2 and Rnd3. The functions of the other members of the Rho GTPase family in cortical development remain largely unknown.

In the Rho subgroup, *RhoA* and *RhoB* are highly expressed in the embryonic cerebral cortex but with distinct patterns (Olenik et al., 1999; Ge et al., 2006; Heng et al., 2008). *RhoA* mRNA is mainly expressed in domains of cellular proliferation whereas *RhoB* mRNA is absent in the proliferative zones but highly expressed in the cortical plate (CP) where neurons migrate or settle at the end of their migration (**Figure 3**). *RhoC* mRNA is detected in the nervous system but its distribution in the developing cerebral cortex has not been thoroughly examined (Erschbamer et al., 2005). Three of the 4 vertebrate Racrelated genes, namely *Rac1*, *Rac3*, and *RhoG*, are expressed in the nervous system (**Figure 3**) (de Curtis, 2008). Most studies

**cortex.** Schematic representation of cortical domains depicting the expression pattern of Rho GTPase genes within the murine cerebral cortex at different developmental stages. To simplify the representation, this figure does not include spatial differences, which should be however kept in mind since all cortical areas do not develop at the same rate and timing. References cited in the Section EXPRESSION OF Rho GTPases IN THE

expression patterns and changes in expression during development. Dark gray and light gray indicate higher and lower relative levels of expression, respectively. VZ: ventricular zone; SVZ: ventricular zone; IZ: intermediate zone; CP: cortical plate; WM: white matter; PP: preplate; n.e.: not expressed in the brain; n.d.: expression in the cerebral cortex not determined.

on cortical neuron development have focused on Rac1, whereas only a few studies exist on Rac3 and RhoG, despite their expression in the developing cerebral cortex. However, their temporal expression is different as illustrated by low levels of *Rac3* mRNA in the embryonic cortex where *Rac1* and *RhoG* mRNA are highly expressed (Ishikawa et al., 2002; Bolis et al., 2003; Corbetta et al., 2005; Fujimoto et al., 2009). Instead, *Rac3* is mainly expressed in the postnatal cortex (P7) in layer V and to a lesser extent in layers II-III (Corbetta et al., 2005). In the Cdc42 subfamily, Cdc42 is expressed throughout the developing cerebral wall (Olenik et al., 1999; Yokota et al., 2010). Cdc42 protein is particularly enriched at the apical/ventricular side of the neuroepithelium and is present in basally located post-mitotic neurons (Cappello et al., 2006). In contrast to Cdc42, *TC10* expression is very low in the embryonic cerebral cortex (**Figure 3**) (Tanabe et al., 2000). Its expression in the brain however increases with development (Tanabe et al., 2000; Abe et al., 2003). Similarly, very low levels of *TCL* mRNA are detected in the brain, at least in adult murine tissue (Vignal et al., 2000). Concerning the last subfamily of classical Rho GTPases, although *RhoF* and *RhoD* are expressed in the adult brain (Murphy et al., 1996; Ellis and Mellor, 2000) and have been shown to regulate neuronal development *in vitro* (Hotulainen et al., 2009; Gad and Aspenstrom, 2010), their expression pattern in the developing cerebral cortex has not been described.

Among the atypical Rho GTPases, the expression profile of Rnd members in the developing cerebral cortex has been the most well-characterized (**Figure 3**). At E14.5, *Rnd1* mRNA levels are very low in the cortex but they gradually increase to peak at postnatal stages (Ishikawa et al., 2003). Conversely, the expression of *Rnd2* and *Rnd3* is high in the embryonic cerebral cortex but show different distribution in the cortical domains (**Figure 3**) (Azzarelli et al., 2014). *RhoV* mRNA is expressed in the human fetal brain (Katoh, 2002) and *RhoU* mRNA is found in the adult human cerebral cortex (Kirikoshi and Katoh, 2002) but their expression in the developing cerebral cortex has not been explored. Like Rac2, the expression of RhoH is restricted to hematopoietic stem cells (Troeger and Williams, 2013).  However, RhoBTB3 is not included in the Rho GTPase family since it does not seem to have a GTP-binding domain, at least it does not contain a consensus GTP-binding motif (Aspenstrom et al., 2007).

#### **Rho GTPases AND REGULATION OF CORTICAL PROJECTION NEURON DEVELOPMENT**

#### **Rho GTPases AND REGULATION OF ADHERENS JUNCTION INTEGRITY**

Like NEs, RGs are highly polarized along their apico-basal axis. They are attached to the luminal surface of the ventricle on their apical side, where they form adherens junctions (AJs) with neighboring RGs (**Figure 4**, ❶), and to the basal lamina via integrins (Gotz and Huttner, 2005). Their cell bodies are retained within the ventricular zone (VZ), a defined region next to the ventricles. AJs between RGs maintain VZ integrity and cortical architecture as well as RG behavior by anchoring a variety of proteins (N-cadherin, β-catenin, αE-catenin) to the actin cytoskeleton (Gotz and Huttner, 2005).

RhoA plays a critical role in the maintenance of these adherens junctional complexes. Indeed, the deletion of *RhoA* by *FoxG1Cre* (Katayama et al., 2011) or *Emx1-Cre* mediated recombination (Cappello et al., 2012) leads to a disorganization of the VZ surface and to a loss of catenin expression at the apical surface around E14.5. In these mutants, rings of intense catenin expression are instead observed inside the brain mass (Katayama et al., 2011; Cappello et al., 2012). Similarly, perturbation of Rho by electroporation either with the RhoA/B/C inhibitor C3 transferase or with *RhoA*, *RhoB*, *RhoC* shRNAs impairs the apical actin filament belt and the apico-basal polarity of electroporated cells (Thumkeo et al., 2011). Expression of dominant-active Rho also affects the actin structure and the apical localization of Ncadherin, suggesting that balanced Rho activity is necessary for maintaining AJ integrity in RGs (Thumkeo et al., 2011) (**Table 1**). While Rho is essential for the maintenance, Cdc42 seems to be essential for the initial formation of apical AJs since the disappearance of apical proteins (PAR6, aPKC, E-cadherin, β-catenin, F-actin and Numb) as well as the apico-basal polarity occurs as early as E10.5 in *FoxG1Cre Cdc42* null embryos (Chen et al., 2006). A similar phenotype is found after deletion of *Cdc42* by *Emx1Cre* or *Nestin-Cre* mediated recombinations (Cappello et al., 2006; Garvalov et al., 2007; Peng et al., 2013) (**Table 1**). Although Rac1 and Cdc42 share many effectors, junction formation and cell polarity during cerebral cortex development specifically require Cdc42 but not Rac1. In Rac1 mutants, VZ progenitors are not tightly packed or radially oriented as in controls. However, this phenotype is not due to an alteration of VZ progenitor polarity since the loss of *Rac1* does not affect the expression pattern of β-catenin and cadherin (Leone et al., 2010). Accordingly, the expression of phosphorylated PAK (p21- activated kinase), which is a direct downstream effector of both Rac1 and Cdc42, at the apical surface of VZ progenitors is affected by the loss of *Cdc42* but not *Rac1* (Leone et al., 2010), further demonstrating that these two Rho GTPases perform non-overlapping and nonredundant functions in the VZ. More recently, the atypical Rho GTPase Rnd3 has also been shown to maintain the integrity of the junctions between RGs through regulation of RhoA and the actin cytoskeleton (Pacary et al., 2013) (**Table 1**).

#### **Rho GTPases AND REGULATION OF INTERKINETIC NUCLEAR MIGRATION**