Neuroscience

The size of the neocortex and its morphology are highly divergent across mammalian species. Several approaches have been utilized for the analysis of neocortical development and comparison among different species. In the present protocol (Note: This protocol requires basic knowledge of brain anatomy), we describe three ex vivo neocortical slice/tissue culture methods: (i) organotypic slice culture (mouse, ferret, human); (ii) hemisphere rotation culture (mouse, ferret); and (iii) free-floating tissue culture (mouse, ferret, human). Each of these three culture methods offers distinct features with regard to the analyses to be performed and can be combined with genetic manipulation by electroporation and treatment with specific inhibitors. These three culture methods are therefore powerful techniques to examine the function of genes involved in neocortical development.

Organotypic slice culture is a powerful technique for exploring the embryonic development of the mammalian brain. In this protocol we describe a basic slice culture technique we have used for two sets of experiments: axon guidance transplant assays and bead culture assays.

Developing axons change responsiveness to guidance cues during the journey to synapse with target cells. Axon crossing at the ventral midline serves as a model for studying how axons accomplish such a switch in their response. Although primary neuron culture has been a versatile technique for elucidating various developmental mechanisms, many in vivo characteristics of neurons, such as long axon-extending abilities and axonal compartments, are not thoroughly preserved. In explant cultures, such properties of differentiated neurons and tissue architecture are maintained. To examine how the midline repellent Slit regulated the distribution of the Robo receptor in spinal cord commissural axons upon midline crossing and whether Robo trafficking machinery was a determinant of midline crossing, novel explant culture systems were developed. We have combined an “open-book” spinal cord explant method with that devised for flat-mount retinae. Here we present our protocol for explant culture of embryonic mouse spinal cords, which allows flexible manipulation of experimental conditions, immunostaining of extending axons and quantitative analysis of individual axons. In addition, we present a modified method that combines ex vivo electroporation and “closed-book” spinal cord explant culture. These culture systems provide new platforms for detailed analysis of axon guidance, by adapting gene knockdown, knockout and genome editing.

This protocol describes the technique of ex-vivo electroporation to target embryonic hippocampal progenitors in an organotypic slice preparation. This technique allows gene perturbation for examining developmental processes in the embryonic hippocampus while retaining the environment and connectivity of the cells. Gene perturbation can include Cre-mediated recombination, RNAi-mediated knockdown, gene overexpression, or a combination of any of these. Ex-vivo electroporation can be performed at a wide range of embryonic stages, giving temporal control to the experimenter. Spatial control can be achieved more easily by preparing the brain in a Petri dish to target particular regions of the hippocampus. The electroporated explant cultures provide a highly tractable system for the study of developmental processes that include progenitor proliferation, migration and cell fate acquisition.

A major issue in developmental biology is to determine how embryonic tissues respond to molecular signals in a timely manner and given the position-restricted instructions during morphogenesis, of which Meckel’s cartilage is a classical example. The ex-vivo explant model is a practical and convenient system that allows investigation of bone and cartilage responses to specific stimuli under a controlled manner that closely mimics the in vivo conditions. In this protocol, the explant model was applied to test whether Meckel’s cartilage and surrounding tissues are responsive to the Endothelin1 (Edn1) signaling molecule and whether it would rescue the phenotype of genetic mutations. The system allows a high degree of manipulation in terms of the concentrations of exogenous compounds added to the explant, time points with regards to measuring mandibular development, and the method of application of exogenous molecules and teratogens.

In the developing and mature central nervous system (CNS) the ventricular lumen is lined by the neuroepithelium and ependymal, respectively. These ventricular epithelia perform important functions related to the development, morphogenesis and physiology of the brain. In the mature CNS, ependyma constitutes a barrier between brain parenchyma and cerebro- spinal fluid (CSF). The most prominent feature of the apical surface of ependymal cells is the presence of multiple motile cilia that extend towards the ventricular lumen. The beating of cilia ensures the circulation of the CSF and its impairment leads to hydrocephalus. For an effective CSF flow, ciliary beating must be coordinated at the level of individual cells and at the tissue level. This coordination is achieved through the precise organization of cilia positioning within the plane of the ependyma. Two major features have been described regarding the planar organization of cilia in ependymal cells (Mirzadeh et al., 2010) and both have a cellular and tissular aspect (Boutin et al., 2014). The first one, rotational polarity, refers to the orientation of ciliary beating. At the cellular level, all cilia beat in the same direction (Figure 1B, black arrows). At the tissue level, each ependymal cell coordinates the direction of their beating with that of neighboring cells (Figure 1C, grey arrows). The second feature, translational polarity, is unique to ependymal cells and refers to the clustering of cilia in a tuft. At the cellular level, this tuft is displaced relative to the center of the ependymal cell (Figure 1B, red arrow). At the tissue level, the positioning of the ciliary tuft is coordinated between adjacent cells (Figure 1C). Alteration of any of these polarities at either level impairs CSF flow circulation (Mirzadeh et al., 2010; Boutin et al., 2014; Guirao et al., 2010; Hirota et al., 2010; Ohata et al., 2014). Cilia axonemes arise from basal bodies (BB) which are cylindrical structures anchored perpendicular to the sub-apical surface of the cells (Figure 1D). BBs are polarized by the presence of appendices such as basal foot or striated rootlets. The basal foot protrudes in a direction correlated with the direction of cilia beating, while the striated rootlet protrudes in the opposite direction of cilia beating (Marshall, 2008). The ‘en face view’ observation of BBs’ organization allows the visualization of ependymal polarities (Mirzadeh et al., 2010; Boutin et al., 2014). Here, we describe an immunofluorescence (IF) protocol for observation of ciliated cells in mouse brain ventricular lateral wall whole mounts (LWWM). This protocol can be used for classical confocal microscopy analysis. In addition, it is well suited for super-resolution STimulated Emission Depletion (STED) microscopy if observation of structures that have features which are smaller than the optical diffraction limit is needed. Finally, we describe a combination of antibodies that allow the concomitant observation, in a single sample, of ependymal polarities at the level of individual cilia, individual cells and at the tissue level.

The neural tube explant culture technique allows in vitro culturing of small pieces of neural tissue isolated from e.g., chick or mouse embryonic tissue in a matrix of collagen for defined periods of time. This method can be used to study the effects of defined molecules on developmental processes such as neural progenitor proliferation and neuronal differentiation and/or survival. Since the explant material can also be prepared from embryonic tissue electroporated with expression vectors, this technique can be adapted to study gene function in the presence of specific environmental signals. Different regions of the neural tube can also be isolated during the dissection step, allowing specific regions of the neural tube to be studied separately. Here, we present a neural tube explant culture method that we have used in several studies (Dias et al., 2014; Lek et al., 2010; Vallstedt et al., 2005).