T&#945;1 promoter is a gift from P. Barker and F.D. Miller. Dcx promoter is a gift from Q. Lu. hGFAP-Cre was a gift from Albee Messing. The PiggyBac transposon vector system was provided by the Sanger Institute. Flt1-DsRed mice were provided by M. Ema (Shiga University). This work was supported by JSPS KAKENHI (Grant Number JP21K07309 to H. Tabata, JP20H05688 and JP22K19365 to K. Nakajima) and Takeda Science Foundation, Keio Gijuku Fukuzawa Memorial Fund for the Advancement of Education and Research, Keio Gijuku Academic Development Funds to K. Nakajima.

The present study was performed with the approval of and following the guidelines of the Animal Care and Use Committee of the Institute for Developmental Research, Aichi Developmental Disability Center (#2019-013), and Keio University (A2021-030).&#160;Timed pregnant ICR (wild-type) mice were obtained commercially (see Table of Materials). To observe the relationship between migrating cells and blood vessels, Flt1-DsRed mice, in which the endothelial cells express DsRed22, were used. Male Flt1-DsRed mice were crossed with female ICR mice (both mice were at the age of 8-24 weeks). All the animals were housed and manipulated under specific pathogen-free (SPF) conditions until sampling. The reagents and equipment used in this study are listed in the Table of Materials.
1. In utero electroporation
<ol>
	<li>Prepare HEPES-buffered saline (HBS) solution by adding an equal volume of autoclaved water to commercially available 2x HBS.</li>
	<li>There are two options of anesthetics; isoflurane (inhalation) and MMB (injection). In the case of using MMB, prepare it as follows. Add 0.75 mL of medetomidine (1 mg/mL), 2 mL of midazolam (5 mg/mL), and 2.5 mL of butorphanol (5 mg/mL) in 19.75 mL of autoclaved water23. 
	NOTE: This anesthetic solution becomes less active within 2 months. Store at 4 &#176;C with light-shield.</li>
	<li>Prepare atipamezole solution. Dilute 0.75 mL of atipamezole (5 mg/mL) with 24.25 mL of autoclaved water.</li>
	<li>Prepare 0.2% (w/v) Fast Green solution in autoclaved water. Sterilize with a filter (pore size 0.22 &#181;m).</li>
	<li>Prepare plasmid using a conventional plasmid purification kit based on alkaline lysis and column purification23 according to the manufacturer&#39;s instructions (see Table of Materials). Dissolve the purified plasmid from 100 mL bacteria culture in 30-50 &#181;L of HBS to prepare the plasmid stock (3-5 &#181;g/&#181;L) (Supplementary File 1).
	<ol>
		<li>Mix and dilute the plasmid stocks with HBS, and add 1/10 volume of 0.2% Fast Green to color the injectant. The components and final concentrations of the plasmids are determined according to their purpose (Table 1).</li>
	</ol>
	</li>
	<li>Make the micropipettes by pulling the glass capillaries using a pipette puller. 
	NOTE: The tip of the pipette is cut obliquely using fine forceps. The outer diameter of the tip is approximately 70 &#181;m. Mark the micropipettes at 2 &#181;L intervals by sucking 2 &#181;L water.</li>
	<li>Inject MMB anesthetic at a dose of 10 &#956;L/g body weight intraperitoneally into a pregnant mouse at an appropriate stage (following institutionally approved protocols). 
	NOTE: The embryonic stages to be manipulated should be determined according to the purpose. To label future II/III pyramidal neurons, E14.5 is suitable. To label glial progenitors, electroporation must be performed on E15-16. Alternatively, the mice can be anesthetized by inhalation of isoflurane (1%-2%).</li>
	<li>After the mouse is deeply anesthetized, place it on a warm plate at 38 &#176;C. 
	NOTE: Ensure that the mouse does not respond to pinching the tail.</li>
	<li>After removing fur around the surgical site using depilatory cream, disinfect the surgical area several times in a circular motion with both an iodine-based scrub and 70% alcohol.&#160;Make a 2 cm long skin incision along the midline from the level of the 2nd most posterior pair of nipples anteriorly, followed by a 2 cm midline incision in the abdominal wall along the linea alba (Figure 1A). 
	NOTE: In the process of incision in the abdominal wall, lift it slightly with fine forceps so as not to cut the internal organs.</li>
	<li>Place a sterile plastic sheet on the skin and a piece of sterile tissue paper on it (Figure 1A). Create a hole in the center to expose the surgical area. 
	NOTE: The sterile plastic sheet and the sterile tissue paper are placed after making the incision because a wide view field is required to position the incision referring to the position of the nipples.</li>
	<li>Wet the paper with sterile PBS, carefully observe the uterine horns from the incision, and draw out the uterine horn through the holes of the plastic sheet and paper.</li>
	<li>Using a micropipette attached to an aspirator tube assembly, inject 1 &#181;L of the plasmid mix into the left or right lateral ventricle through the uterine wall by expiratory pressure (Figure 1A). 
	NOTE: Alternatively, plasmid can be injected by using a microinjector.</li>
	<li>Pinch the head of the embryo with a tweezer-type electrode and apply electronic pulses using an electroporator (35 V for E14-16, 50 V for E17, 50 ms square pulses with 450 ms intervals, 5 times) (Figure 1A). 
	NOTE: Do not pinch the head of the embryos strongly, which will damage the embryos. The actual current should be 40-70 mA.</li>
	<li>Place the uterine horn back into the abdominal cavity.</li>
	<li>Suture the abdominal wall using a 5-0 silk thread.</li>
	<li>Apply atipamezole solution at a dose of 10 &#181;L/g body weight intraperitoneally.</li>
	<li>Suture the skin using a 5-0 silk thread. 
	NOTE: The skin can be closed by using an autoclip.</li>
	<li>Keep the manipulated mice on a warm plate for at least 30 min. The mice will start moving in the cage. 
	NOTE: The procedure of in utero electroporation (from steps 1.7 to 1.16) should be done within 30 min. The typical operation time is 20 min.</li>
</ol>
2. Preparation of slices
NOTE: The freshness of the slices is the key to the success of this experiment. The slices must be prepared (from steps 2.5-2.12) within 2 h.
<ol>
	<li>Prepare the Hanks&#39; balanced salt solution with Ca2+ and Mg2+; HBSS(+). Add 5 mL of CaCl2 (14 g/L, autoclaved) and 5 mL of MgSO4.7H2O (20 g/L, autoclaved) to 500 mL of HBSS(-). Store at 4 &#176;C. Bubble the solution with 95% O2 + 5% CO2 gas for 5 min on ice before use.</li>
	<li>Prepare the culture medium. At least 2.5 mL/dish is needed. Add 2 mL of fetal bovine serum, 25 &#181;L of L-glutamine (200 mM), 200 &#181;L B27 (50x), 50 &#181;L of penicillin + streptomycin (10 K units/mL and 10 mg/mL) to 8 mL of Neurobasal medium.</li>
	<li>Prepare 3% low melting temperature agarose gel. Pore 50 mL of HBSS(+), then add 1.5 mg of low melting temperature agarose, and autoclave it. Melt the agarose in a microwave oven and keep it at 50 &#176;C in a water bath incubator prior to use.</li>
	<li>Add 1.9 mL of culture medium to a glass base dish. Put a cell culture insert on it carefully to not trap air bulbs under the filter, then add 500 &#181;L of culture medium onto the filter (Figure 1B). Place it on ice.</li>
	<li>After anesthetizing the manipulated mice by inhalation of 5% isoflurane, sacrifice them by cervical dislocation (following institutionally approved protocols) at appropriate stages to be observed. 
	NOTE: Two days (48 h) and 3 days after electroporation at E14.5 are suitable for observing the multipolar-bipolar transition and locomotion mode, respectively.</li>
	<li>Take out the embryos and place them in a Petri dish with cold PBS.</li>
	<li>Select the embryos by the expression of GFP or other fluorescent proteins under a microscope.</li>
	<li>After decapitation of the embryos, dissect out the brains6 under a dissecting microscope and keep them in cold HBSS(+).</li>
	<li>Pour the melted 3% low melting temperature agarose gel into a cryomold (see Table of Materials), and then put the brains in it and roll them several times. Place it at room temperature until the agarose completely solidifies, and then place it on ice.</li>
	<li>Slice the brains coronally at the thickness of 350 &#181;m using a vibrating microtome (see Table of Materials).</li>
	<li>Arrange the slices on the filter (maximum 6 slices per filter).</li>
	<li>Withdraw the culture medium from the inside and outside of the filter cup (600 &#181;L in total). In this step, the slices are pressed and fixed onto a filter. After 5 min, add 100 &#181;L of culture medium to the inside of the cell culture insert.</li>
</ol>
3. Time-lapse imaging
<ol>
	<li>Keep the culture chamber mounted on the stage of an inverted laser scan microscope at 37 &#176;C using the heater of the chamber or a hood enclosing the microscope at least 30 min before imaging to prevent Z drift. Supply humidified 5% CO2 + 40% O2 gas to the culture chamber.</li>
	<li>Set the culture dish with slices prepared above in the culture chamber.</li>
	<li>Using a long working distance 20x lens (NA = 0.45), acquire around 10 Z-stack images with 5 &#181;m steps every 15-30 min for 24 h. 
	NOTE: The use of a motorized XY stage is highly recommended. This enables us to obtain images from multiple objective fields. Typically, we obtain 10-20 images from different objective fields in one time-lapse experiment.</li>
</ol>
4. Imaging analysis
<ol>
	<li>Convert XYZT images to XYT by the maximum projection of the Z axis using ImageJ or other software. 
	NOTE: If the slices slid during the observation, adjust the position of the cells within the images using StackRed, an ImageJ plugin, with its function of Rigid Body transformation (see Table of Materials).</li>
	<li>Trace the trajectory of GFP- or other fluorescent protein-positive cells either manually or automatically. Manual and automatic tracing can be performed using MTrackJ24 and TrackMate25,26 (ImageJ plugins), respectively. Automatic tracing using TrackMate is efficient for analyzing many cells in an unbiased manner. However, manual tracing using MTrackJ remains useful for the accurate tracking of individual cells.</li>
</ol>

Visualizing Neuronal and Glial Cell Dynamics in the Developing Mouse Brain

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+hyperactive+piggyBac+transposase+for+mammalian+applications.">A hyperactive piggyBac transposase for mammalian applications.</a> P Natl Acad Sci USA. 108 (4), 1531-1536 (2011).</li><li>Chen, F., LoTurco, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+stable+transgenesis+of+radial+glia+lineage+in+rat+neocortex+by+piggyBac+mediated+transposition.">A method for stable transgenesis of radial glia lineage in rat neocortex by piggyBac mediated transposition.</a> J Neurosci Meth. 207 (2), 172-180 (2012).</li><li>Wang, X., Qiu, R., Tsark, W., Lu, Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+promoter+analysis+in+developing+mouse+brain+and+genetic+labeling+of+young+neurons+by+doublecortin-DsRed-express.">Rapid promoter analysis in developing mouse brain and genetic labeling of young neurons by doublecortin-DsRed-express.</a> J Neurosci Res. 85 (16), 3567-3573 (2007).</li><li>Sauer, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DNA+recombination+with+a+heterospecific+Cre+homolog+identified+from+comparison+of+the+pac-c1+regions+of+P1-related+phages.">DNA recombination with a heterospecific Cre homolog identified from comparison of the pac-c1 regions of P1-related phages.</a> Nucleic Acids Res. 32 (20), 6086-6095 (2004).</li><li>Hermann, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Binary+recombinase+systems+for+high-resolution+conditional+mutagenesis.">Binary recombinase systems for high-resolution conditional mutagenesis.</a> Nucleic Acids Res. 42 (6), 3894-3907 (2014).</li><li>Kanatani, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+COUP-TFII/Neuropilin-2+is+a+molecular+switch+steering+diencephalon-derived+GABAergic+neurons+in+the+developing+mouse+brain.">The COUP-TFII/Neuropilin-2 is a molecular switch steering diencephalon-derived GABAergic neurons in the developing mouse brain.</a> P Natl Acad Sci USA. 112 (36), E4985-E4994 (2015).</li><li>Yozu, M., Tabata, H., Nakajima, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+caudal+migratory+stream:+A+novel+migratory+stream+of+interneurons+derived+from+the+caudal+ganglionic+eminence+in+the+developing+mouse+forebrain.">The caudal migratory stream: A novel migratory stream of interneurons derived from the caudal ganglionic eminence in the developing mouse forebrain.</a> J Neurosci. 25 (31), 7268-7277 (2005).</li><li>Kanatani, S., Yozu, M., Tabata, H., Nakajima, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=COUP-TFII+is+preferentially+expressed+in+the+caudal+ganglionic+eminence+and+is+involved+in+the+caudal+migratory+stream.">COUP-TFII is preferentially expressed in the caudal ganglionic eminence and is involved in the caudal migratory stream.</a> J Neurosci. 28 (50), 13582-13591 (2008).</li><li>Kitazawa, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hippocampal+pyramidal+neurons+switch+from+a+multipolar+migration+mode+to+a+novel+&amp;#34;climbing&amp;#34;+migration+mode+during+development.">Hippocampal pyramidal neurons switch from a multipolar migration mode to a novel &amp;#34;climbing&amp;#34; migration mode during development.</a> J Neurosci. 34 (4), 1115-1126 (2014).</li></ol>

The authors have no conflicts of interest.

This protocol introduced a method for the time-lapse observation of cells derived from the pallial (cortical) VZ. To label the migrating cells from the VZ, we used in utero electroporation, in which individual cells were clearly labeled with a higher signal-to-noise ratio than in viral vector-mediated labeling. Using in utero electroporation, any type of vector in any combination can be easily introduced into the radial glial cells (neural stem cells) in living embryos. Neurons and glial progenitors can be selectively labeled using different promoters. Since the copy number of plasmids introduced by in utero electroporation is high, gain-of-function and loss-of-function studies can be easily conducted by co-electroporation of GFP expression vectors and various vectors related to the genes of interest, such as expression vectors and knockdown/knockout vectors. Moreover, in utero electroporation can be used to introduce plasmid vectors into the VZ of various brain regions other than the cortical VZ, which enables us to observe the migration of other cell types such as inhibitory neurons, oligodendrocyte progenitors from ganglionic eminences34,35,36 and hippocampal pyramidal neurons37.
In time-lapse studies, the effects of gain-of-function and loss-of-function experiments on cell behavior can be evaluated using the TrackMate and MTrackJ software. Many genes, including those responsible for neurodevelopmental disorders and intellectual disabilities, are involved in neuronal migration. Time-lapse studies can directly reveal how these genes regulate cell behavior. Compared with neuronal migration, the molecular mechanisms of glial cell migration are not well understood. This method may reveal not only the normal development of glial cells, but also the genetic and environmental factors that can affect glial development and its relationship with mental illness.
The limitation of this method is that it inevitably depends on the in vitro slice culture method. Interestingly, migrating neurons do not lose directional cues for migration toward the brain surface, even in slices cultivated in vitro. However, investigators must be careful about artifacts. Cells near the surface are attached to the filter, and the edges of the slices are easily deformed. These cells should be excluded from the analysis. Control experiments should be considered if possible. It is important to note that brain slices recapitulate normal development within approximately 24 h. Therefore, the timing of slice preparation from electroporation should be determined so that the events to be analyzed will occur in this time window. To observe the multipolar-bipolar transition, slices should be prepared 2 days after electroporation. To analyze migration in the locomotion mode, 3 days later is appropriate. The observation of blood vessel-guided migration is a more difficult task than that of other experiments. The endothelial cells are easily degraded in slices without blood flow. To minimize blood vessel degradation, the slices should be prepared as quickly as possible and maintained in a medium containing 20% serum and 40% O2 gas. Other complementary experiments, such as histological analyses of fixed brains or in vivo live imaging using two-photon microscopy, may strengthen the observations found in slice time-lapse analyses.
In summary, this method is the only option for observing cell behavior for 24 h. With the development of novel plasmid vectors, the range of applications of this method will expand further in the future.

During the development of the cerebral cortex, (apical) radial glia in the pallial ventricular zone (VZ) lining the lateral ventricle produce first neurons and then glial progenitors with some overlapping period1. Neurons are also generated from intermediate progenitors or basal radial glia in the subventricular zone (SVZ) adjacent to the VZ, both of which originate from the (apical) radial glia2,3. In mice, radial glial cells produce only neurons on embryonic day (E) 12-14, both neurons and glial progenitors on E15-16, and glial progenitors from E17 onward4. The major population of glial progenitors generated during these embryonic stages preferentially differentiates into astrocytes, although some cells also differentiate into oligodendrocytes5. Neurons and astrocyte progenitors generated at these stages migrate toward the brain surface and enter the cortical plate (future cortical gray matter). Neuronal migration from the VZ to the cortical plate occurs in multiple phases. Neurons first adopt a multipolar morphology just above the multipolar cell accumulation zone (MAZ), overlapping the SVZ or intermediate zone, where they vigorously extend and retract multiple thin processes and slowly migrate (multipolar migration)6,7. After approximately 24 h, neurons transform into a bipolar morphology, extending a thick leading process toward the brain surface and a thin trailing process backward, and migrate linearly toward the brain surface using a radial process extending from the radial glia to the pial surface as a scaffold, which is called locomotion mode2,8. Because neurons in locomotion mode always reach the outermost surface of the cortical plate, passing through their predecessors just under the marginal zone, neurons are aligned in a birthdate-dependent inside-out manner in the cortical plate9,10,11.
In contrast, astrocyte progenitors migrate rapidly to the intermediate zone and cortical plate, with frequent directional changes. This migratory behavior is completely different from neuronal migration and is called erratic migration5. Astrocyte progenitors also migrate along blood vessels in a process called blood vessel-guided migration. Astrocyte progenitors switch between these migration modes and reach the cortical plate5,12. Although the positioning of astrocytes is not strictly determined by their date of production, a mild tendency for early-born astrocytes to settle in the superficial part of the cortical plate has been observed5. Interestingly, astrocytes that settle in the cortical plate are generated in embryonic stages and eventually differentiate into protoplasmic astrocytes, whereas postnatally generated astrocytes do not migrate actively, remain in the white matter, and differentiate into fibrous astrocytes5. How this stage-dependent specification of astrocytic subtypes occurs remains unclear.
A growing number of genes involved in neuronal migration have been identified, including those involved in neurodevelopmental and psychiatric disorders13,14. Therefore, it is crucial to elucidate the effects of mutations in these genes on the behavior of migrating neurons. As previously mentioned, neuronal migration occurs in multiple phases. Time-lapse observations can directly determine the phase that is mainly affected (cell cycle exit, multipolar-bipolar transition, migration speed of locomotion, etc.). However, the molecular mechanisms underlying the specification, migration, and positioning of astrocytes remain largely unknown. Given that astrocytes play crucial roles in synaptogenesis15 and blood-brain barrier formation during brain development16, developmental defects in astrocytes may result in neurodevelopmental disorders. Time-lapse studies on astrocyte progenitors may clarify these molecular mechanisms and their relationship with mental illness.
This protocol provides a method for time-lapse observation of cortical VZ-derived cells. A similar video protocol for the observation of neuronal migration has already been published17. Here, we describe the method for both migrating neurons and astrocyte progenitors. To label these cells with fluorescent proteins, such as green and red fluorescent proteins (GFP and RFP), plasmid mixtures containing appropriate components are introduced into the cortical VZ by in utero electroporation at appropriate stages18,19,20,21. The manipulated embryos are removed at the desired stages, and the brains are sliced and used for time-lapse observations using a laser scanning microscope. The migration speed, direction, and other behaviors, which are never addressed using fixed brain samples, can be examined using this method. Using in utero electroporation, expression, and knockdown/knockout vectors can be easily transferred concomitantly with fluorescent protein vectors, enabling us to conduct gain-of-function and loss-of-function studies of specific genes.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Aspirator tube assembly</td>
			<td>Drummond</td>
			<td>2-040-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Atipamezole (5 mg/mL)</td>
			<td>Meiji</td>
			<td>Mepatia</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclip</td>
			<td>Becton Dickinson</td>
			<td>427630</td>
			<td>9 mm</td>
		</tr>
		<tr>
			<td>B27 supplement</td>
			<td>Gibco</td>
			<td>17504-044</td>
			<td></td>
		</tr>
		<tr>
			<td>Butorphanol (5 mg/mL)</td>
			<td>Meiji</td>
			<td>Vetorphale</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture insert</td>
			<td>Millipore</td>
			<td>PICM ORG 50</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Nikon</td>
			<td>A1RHD25</td>
			<td>Equipped with a long working distance lens (S Plan Fluor ELWD 20XC)</td>
		</tr>
		<tr>
			<td>Cryomold</td>
			<td>Tissue-Tek</td>
			<td>4566</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture chamber</td>
			<td>Tokken</td>
			<td>TK-NBCMP</td>
			<td>Custom-made</td>
		</tr>
		<tr>
			<td>Electroporator</td>
			<td>NEPA Gene</td>
			<td>NEPA21</td>
			<td></td>
		</tr>
		<tr>
			<td>Fast Green</td>
			<td>Sigma-Aldrich</td>
			<td>F7258</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas mixer</td>
			<td>Tokken</td>
			<td>TK-MIGM01-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass base dish</td>
			<td>Iwaki</td>
			<td>3910-035</td>
			<td>Diameter of glass base is 27 mm</td>
		</tr>
		<tr>
			<td>Glass capillaries</td>
			<td>Narishige</td>
			<td>GD-1</td>
			<td></td>
		</tr>
		<tr>
			<td>HBS (2x)</td>
			<td>Sigma-Aldrich</td>
			<td>51558</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS(-)</td>
			<td>Wako</td>
			<td>084-08345</td>
			<td></td>
		</tr>
		<tr>
			<td>Heater Unit</td>
			<td>Tokken</td>
			<td>TK-0003HU20</td>
			<td>Custom-made, including hood and heater</td>
		</tr>
		<tr>
			<td>hGFAP-Cre</td>
			<td>Addgene</td>
			<td>#40591</td>
			<td>A gift from Albee Messing</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td></td>
			<td></td>
			<td>https://imagej.net/ij/</td>
		</tr>
		<tr>
			<td>L-glutamine (200 mM)</td>
			<td>Gibco</td>
			<td>25030</td>
			<td></td>
		</tr>
		<tr>
			<td>Low melting temperature agarose</td>
			<td>Lonza</td>
			<td>50100</td>
			<td></td>
		</tr>
		<tr>
			<td>Medetomidine (1 mg/mL)</td>
			<td>Meiji</td>
			<td>Medetomin</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>Narishige</td>
			<td>IM-300</td>
			<td></td>
		</tr>
		<tr>
			<td>Midazolam (5 mg/mL)</td>
			<td>Sandoz</td>
			<td>Midazolam</td>
			<td></td>
		</tr>
		<tr>
			<td>MTrackJ</td>
			<td></td>
			<td></td>
			<td>https://imagescience.org/meijering/software/mtrackj/</td>
		</tr>
		<tr>
			<td>Neurobasal medium</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>pCAG-hyPBase</td>
			<td></td>
			<td></td>
			<td>The hyPBase cDNA from pCMV-hyPBase (a gift from Sanger Institute) was inserted into the downstream of the CAG promoter of pCAGGS (a gift from J. Miyazaki).</td>
		</tr>
		<tr>
			<td>pDcx-Dre</td>
			<td></td>
			<td></td>
			<td>The Dcx promoter from Dcx4kbEGFP70 (a gift from Q. Lu) was exchanged with CAG promoter of pCAG-NLS-HA-Dre34 (a gift from Pawel Pelczar, Addgene #51272).</td>
		</tr>
		<tr>
			<td>Penicillin + Streptomycin</td>
			<td>Gibco</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasmid purification kit</td>
			<td>Invitrogen</td>
			<td>PureLink HiPure plasmid midiprep kit (K210005)</td>
			<td></td>
		</tr>
		<tr>
			<td>pPB-CAG-LNL-RFP</td>
			<td></td>
			<td></td>
			<td>CAG-LNL cassette from pCALNL-DsRed (a gift from Connie Cepko, Addgene #13769), and TurboRFP cDNA (Evrogen, FP232) were inserted into the cloning site of pPB-CAG.EBNXN (a gift from Sanger Institute).</td>
		</tr>
		<tr>
			<td>pPB-CAG-rDIO-EGFP</td>
			<td></td>
			<td></td>
			<td>The sequence containning synthetic rox sites, synthetic DIO cassette, and EGFP cDNA from pEGFP-N1 (Clontech, U55762) in reverse direction&#160; were inserted into the cloning site of pPB-CAG.EBNXN (a gift from Sanger Institute). The sequence is provided in the Supplementary File.</td>
		</tr>
		<tr>
			<td>Puller</td>
			<td>Narishige</td>
			<td>PN-31</td>
			<td></td>
		</tr>
		<tr>
			<td>StackRed</td>
			<td></td>
			<td>a plugin for ImageJ</td>
			<td>http://bigwww.epfl.ch/thevenaz/stackreg/</td>
		</tr>
		<tr>
			<td>Suture needle</td>
			<td>Nazme</td>
			<td>C-24-521-R No.1</td>
			<td>1/2 circle, length 14 mm</td>
		</tr>
		<tr>
			<td>Suture thread</td>
			<td>Nazme</td>
			<td>C-23-B2</td>
			<td>Silk, size 5-0</td>
		</tr>
		<tr>
			<td>Timed pregnant ICR (wild-type) mice</td>
			<td>Japan SLC</td>
			<td>ICR mouse</td>
			<td></td>
		</tr>
		<tr>
			<td>TrackMate</td>
			<td></td>
			<td></td>
			<td>https://imagej.net/plugins/trackmate/index</td>
		</tr>
		<tr>
			<td>Tweezer-type electrode</td>
			<td>BEX or NEPA Gene</td>
			<td>CUY650P5&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>T&#945;1-EGFP</td>
			<td></td>
			<td></td>
			<td>EGFP cDNA from pEGFP-N1 (Clontech, U55762) was inserted into the downstream of the T&#945;1 promoter in plasmid 253 (a gift from P. Barker and F.D.Miller)</td>
		</tr>
		<tr>
			<td>Vibrating microtome</td>
			<td>Leica or Zeiss</td>
			<td>Vibrating blade microtome VT1000S or Hyrax V50.</td>
			<td></td>
		</tr>
	</tbody>
</table>

time-lapse imaging of migrating neurons and glial progenitors in embryonic mouse brain slices

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain surface. This process is crucial for proper brain function, and its dysregulation can result in neurodevelopmental and psychiatric disorders after birth. In fact, many genes responsible for these diseases have been found to be involved in this process, and therefore, revealing how these mutations affect cellular dynamics is important for understanding the pathogenesis of these diseases. This protocol introduces a technique for time-lapse imaging of migrating neurons and glial progenitors in brain slices obtained from mouse embryos. Cells are labeled with fluorescent proteins using in utero electroporation, which visualizes individual cells migrating from the ventricular zone with a high signal-to-noise ratio. Moreover, this in vivo gene transfer system enables us to easily perform gain-of-function or loss-of-function experiments on the given genes by co-electroporation of their expression or knockdown/knockout vectors. Using this protocol, the migratory behavior and migration speed of individual cells, information that is never obtained from fixed brains, can be analyzed.

Radial glial cells in the pallial VZ produce only neurons until E14, and both neurons and glial cells at E15 and E16. To observe the migratory behaviors of neurons and glial cells simultaneously, we labeled them with enhanced GFP (EGFP) and RFP, respectively, by using a neuron-specific promoter, T&#945;1 promoter27, and human glial fibrillary acidic protein (hGFAP) promoter28, which is preferentially activated in astrocytes. Astrocyte progenitors repeat cell divisions during the migration, leading to the dilution of plasmid in the cells. To avoid this, a transposon vector system consisting of a donor plasmid (pPB) and a transposase expression vector (pCAG-hyPBase) was used29,30. The donor plasmid used here (pPB-CAG-LNL-RFP) has loxP-neo-loxP (LNL) cassette, and expresses RFP depending on Cre recombinase. ICR mice (wild type; WT) were electroporated with the indicated plasmids in Figure 2A at E15, and the slices were prepared 3 days later. During this period of time, the first cohort of EGFP+ neurons reached the surface of the cortical plate (Figure 2B), but many EGFP+ neurons were still migrating. Time-lapse observation was performed for 24 h. During this observation, neurons with a thick leading process migrated linearly toward the brain surface at a relatively constant rate in the cortical plate (19.54 &#177; 6.21 &#181;m/h, mean &#177; standard deviation, 70 cells). The migration trajectories were traced automatically by using TrackMate software (magenta lines in Figure 2B). In the same objective field, astrocyte progenitors were visualized with RFP (Figure 2C). Because the hGFAP promoter is also active in radial glial cells in late embryonic stages, some neurons derived from those radial glia also express RFP (blue arrows in Figure 2B,C). These neurons are almost always transfected with an EGFP expression vector. Therefore, astrocyte progenitors were identified as RFP single-positive cells.&#160;
In contrast to neurons, astrocyte progenitors migrate rapidly and randomly with frequent directional changes within the intermediate zone and cortical plate (erratic migration)5. Migration of astrocyte progenitors was traced manually by MTrackJ software (magenta lines in Figure 2C), which revealed their high motility (88.37 &#177; 33.61 &#181;m/h, 5 cells). To observe the relationship between astrocyte progenitors and blood vessels, Flt1-DsRed mice, which express DsRed, an RFP, in endothelial cells, were used. In this experiment, the astrocyte progenitors were supposed to be identified only by EGFP. To this end, the rDIO system5 was developed, in which EGFP expression depending on hGFAP-Cre can be eliminated in neurons by using a Dcx promoter (an early neuron-specific promoter)31-dependent activation of another set of DNA recombinase and its recognition sequence (Dre-rox) (Figure 2D, the sequence of pPB-CAG-rDIO-EGFP is provided in Supplementary File 1)32,33. The rDIO system was introduced into Flt1-DsRed mouse at E15 by in utero electroporation, and slices were prepared 24 h later. As a result, it was observed that the EGFP-positive astrocyte progenitors actively migrated along blood vessels (blood vessel-guided migration)5 (Figure 2E).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66631/66631fig01.jpg" /> 
Figure 1: Flow of time-lapse observations of cells labeled by in utero electroporation. (A) A pregnant mouse at an appropriate stage is anesthetized, and the abdominal cavity is cut open. The surgical area is covered with sterile plastic and sterile paper sheets. The uterine horns are exposed, and embryos are injected with 1 &#181;L of plasmid mixture and applied electronic pulses. The abdominal wall and skin are sutured, and the manipulated mice are placed back in the home cage. (B) Slices prepared from electroporated brains are set on the filter of cell culture insert (6 slices maximum) in a glass base dish. The glass base dishes containing slices are mounted in a culture chamber. This time-lapse system has an acryl hood covering the whole microscope to keep the temperature at 37 &#176;C. The culture chamber, in which humidified mixed gas is supplied, is equipped with a microscope stage inside the hood. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66631/66631fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66631/66631fig02.jpg" /> 
Figure 2: Time-lapse observations of neurons and astrocyte progenitors. (A) Method for differential labeling of neurons (EGFP, green) and astrocyte progenitors (RFP, red), and a representative image of a slice prepared 2 days after the electroporation at 2 h from the start of time-lapse imaging (right). (B,C) Time-lapse images of EGFP+ (B) and RFP+ cells (C) on the same objective field shown in (A)-right from 2 h to 21 h of observation. (B) Trajectories of EGFP+ migrating cells detected automatically by TrackMate software were shown in magenta. (C) Trajectories of RFP+ cells were traced manually using MTrackJ software. EGFP/RFP double positive cells, which are thought to be neurons, are indicated by blue arrows. (D) Labeling strategy of astrocyte progenitors using rDIO system in Flt1-DsRed mouse embryos. (E) Representative result of blood vessel-guided migration. A cell (arrow) protruded a leading process (arrowhead), and migrated along the DsRed-expressing blood vessel (magenta) toward the pial surface. CP; cortical plate. IZ; intermediate zone. VZ; ventricular zone. Scale bars: 200 &#181;m (A&#8211;C), 50 &#181;m (E). This figure is adapted from Tabata H. et al.5. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66631/66631fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Neurons (layers II/III)</td>
			<td colspan="2">Astrocyte progenitors</td>
			<td colspan="2">rDIO system 
			(astrocyte specific labeling)</td>
		</tr>
		<tr>
			<td>pCAG-EGFP</td>
			<td>0.5 &#181;g/&#181;L</td>
			<td>pPB-CAG-LNL-EGFP</td>
			<td>0.5 &#181;g/&#181;L</td>
			<td>pPB-CAG-rDIO-EGFP</td>
			<td>0.5 &#181;g/&#181;L</td>
		</tr>
		<tr>
			<td>or Ta1-EGFP (more specific for neurons)</td>
			<td>0.5 &#181;g/&#181;L</td>
			<td>hGFAP-Cre</td>
			<td>0.75 &#181;g/&#181;L</td>
			<td>hGFAP-Cre</td>
			<td>0.75 &#181;g/&#181;L</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>pCAG-hyPBase</td>
			<td>0.5 &#181;g/&#181;L</td>
			<td>pDcx-Dre</td>
			<td>0.5 &#181;g/&#181;L</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>pCAG-hyPBase</td>
			<td>0.5 &#181;g/&#181;L</td>
		</tr>
	</tbody>
</table>
Table 1: Representative components of plasmid mix for each target. EGFP vectors can be replaced with equivalent RFP or other fluorescent protein expression vectors. To label astrocyte progenitors, a transposon vector system is used. pPB vectors are PiggyBac transposon vectors for genome integration. pCAG-hyPBase is a PiggyBac transposase expression vector. LNL is a loxP-neo-loxP cassette for Cre-dependent expression. rDIO-EGFP is Cre-loxP and Dre-roxP dependent on/off cassette of EGFP expression. To observe the gain-of-function or loss-of-function phenotype of desired genes, expression vectors or knockdown vectors can be added at 1 &#181;g/&#181;L. The concentration of these vectors should be adjusted in each case.
Supplementary File 1: Plasmid sequences. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66631/Supplementary_File.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Author Spotlight: Exploring Cell Migration and Gene Roles in the Developing Brain at JoVE.com

Author Spotlight: Exploring Cell Migration and Gene Roles in the Developing Brain

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain surface. Many genes are involved in this process. This protocol introduces the technique for the time-lapse imaging of migrating neurons and glial progenitors.

Time-Lapse Imaging of Migrating Neurons and Glial Progenitors in Embryonic Mouse Brain Slices

During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain ...

time-lapse-imaging-migrating-neurons-glial-progenitors-embryonic

Nanyang Technological University

Research

JoVE Journal

Neuroscience

740 Views.  Keio University School of Medicine. During the development of the cerebral cortex, neurons and glial cells originate in the ventricular zone lining the ventricle and migrate toward the brain surface. Many genes are involved in this process. This protocol introduces the technique for the time-lapse imaging of migrating neurons and glial progenitors.

Video: Author Spotlight: Exploring Cell Migration and Gene Roles in the Developing Brain

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as well as differentiation and integration of new neurons into existing neural circuits. For postnatal neurogenesis in the olfactory bulbs, these events are segregated within three anatomically distinct domains: proliferation largely occurs in the subependymal zone (SEZ) of the lateral ventricles, migrating neuroblasts traverse through the rostral migratory stream (RMS), and new neurons differentiate and integrate within the olfactory bulbs (OB). The three domains serve as ideal platforms to study the cellular, molecular, and physiological mechanisms that regulate each of the biological events distinctly. This paper describes an organotypic slice assay optimized for postnatal brain tissue, in which the extracellular conditions closely mimic the in vivo environment for migrating neuroblasts. We show that our assay provides for uniform, oriented, and speedy movement of neuroblasts within the RMS. This assay will be highly suitable for the study of cell autonomous and non-autonomous regulation of neuronal migration by utilizing cross-transplantation approaches from mice on different genetic backgrounds.

This protocol describes an organotypic slice assay optimized for the postnatal brain and high-resolution time-lapse imaging of neuroblast migration in the rostral migratory stream.

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Efficient Derivation of Human Neuronal Progenitors and Neurons from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system (CNS) structure and circuitry in today's healthcare industry. Cell-based therapies hold great promise to restore the lost nerve tissue and function for CNS disorders. However, cell therapies based on CNS-derived neural stem cells have encountered supply restriction and difficulty to use in the clinical setting due to their limited expansion ability in culture and failing plasticity after extensive passaging1-3. Despite some beneficial outcomes, the CNS-derived human neural stem cells (hNSCs) appear to exert their therapeutic effects primarily by their non-neuronal progenies through producing trophic and neuroprotective molecules to rescue the endogenous cells1-3. Alternatively, pluripotent human embryonic stem cells (hESCs) proffer cures for a wide range of neurological disorders by supplying the diversity of human neuronal cell types in the developing CNS for regeneration1,4-7. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity7-10. In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic11-13. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules14 (please see a schematic in Fig. 1). Retinoic acid (RA) does not induce neuronal differentiation of undifferentiated hESCs maintained on feeders1, 14. And unlike mouse ESCs, treating hESC-differentiated embryoid bodies (EBs) only slightly increases the low yield of neurons1, 14, 15. However, after screening a variety of small molecules and growth factors, we found that such defined conditions rendered retinoic acid (RA) sufficient to induce the specification of neuroectoderm direct from pluripotent hESCs that further progressed to neuroblasts that generated human neuronal progenitors and neurons in the developing CNS with high efficiency (Fig. 2). We defined conditions for induction of neuroblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human neuronal cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of neuroblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human neuronal progenitors and neuronal cell types in the developing CNS for neural repair.

There is a large unfulfilled need for a clinically-suitable human neuronal cell source for repair or regeneration of the damaged central nervous system ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...