M.S. and Y.Z. are supported by KCL and KCL-China PhD studentships. M.O. was funded by a Biotechnology and Biological Sciences Research Council PhD studentship. We thank Masaru Okabe and Jun-ichi Miyazaki for the pCX-EGFP plasmid and Alain Chedotal and Athena Ypsilanti for valuable advice on electroporation.

This procedure is in accordance with the UK Home Office Regulations (Animal Scientific Procedures Act, 1986). Scientists should follow the guidelines established and approved by their institutional and national animal regulatory organizations.
1. Postnatal Electroporation
1.1. Preparation of Glass Capillaries, DNA Solution and Electroporator
<ol>
	<li>Prepare pulled glass capillaries (O.D.: 1.5 mm, I.D.: 0.86 mm) for DNA injection. (Indicative settings for a Sutter P-97 capillary puller are: Heat 283; Pull 50; Velocity 90; Time 50). Make a mark on the capillary corresponding to a volume of approximately 2 &#181;l.</li>
	<li>Set the voltage of the electroporator to 5 square pulses, 50 msec/pulse at 100 V, with 850 msec intervals (100 V, pulse ON 50 msec, pulse OFF 850 msec, pulse 5).</li>
	<li>Dilute high-purity (OD 260/280 &#62; 1.80) endotoxin-free plasmid DNA to a final concentration of 1-2 &#181;g/&#181;l with endotoxin free Tris-EDTA buffer or PBS. It is recommended to add 0.1% Fast Green to the DNA solution (the dye should spread in the ventricle when successfully injected).</li>
	<li>Prepare the 5-7 mm electrodes, the electrode gel and warm up the heating pad.</li>
</ol>
1.2. Electroporation
<ol>
	<li>Remove a postnatal day 2 mouse pup from cage and anesthetize it by isoflurane inhalation (at a flow of ~0.6 L/min).</li>
	<li>After approximately 1 min, determine the state of anesthetization using the foot pinch response. If no movement occurs, proceed with intraventricular injection.</li>
	<li>Load capillary needle with 1-2 &#181;l of DNA using an aspirator tube connected to the capillary.</li>
	<li>Under a cold light source, hold the head of the pup between your thumb and index finger of your less dominant hand. Slightly pull the skin back on the head to help identification of the right injection point.</li>
	<li>Consider a virtual line between the eye and the craniometric landmark lambda (Figure 1A). Insert the capillary needle at about one third of the length of this line from the lambda (about 1 mm from the line midpoint)15. Insert the capillary about 2 mm deep, making sure to avoid deep penetration in the brain.</li>
	<li>Inject plasmid by blowing slowly through mouth (a syringe connected to the capillary can also be used). During this procedure, make sure your fingers are not applying too much pressure on the brain, as this can prevent successful plasmid injection.</li>
	<li>Stop injection when a minimal amount of DNA solution is left in the capillary. It is advisable to inject less than 1 &#181;l to avoid deleterious increase in intracranial pressure.</li>
	<li>Coat both electrodes with gel and place them with the positive side on the lateral side of the hemisphere where the DNA was injected (Figure 1A). For DNA incorporation into the rostral SVZ, place electrodes slightly rostral to the injection point. Varying electrode position can achieve regional specificity of electroporation in different areas of the SVZ22,25.</li>
	<li>Initiate current transfer by pressing the pulse footswitch pedal. When the electroporation is complete, check the voltage on the electroporator display (voltage values should not be below 90 V, since lower voltage&#160;values correlate with poor electroporation efficiency).</li>
	<li>Reanimate the pup under oxygen on heating pad for a few minutes and return it to the cage, placing it away from the mother. Make sure the mother retrieves the pup and reunites it with the rest of the litter. After electroporation, leave pups with their mother for 4-7 days before next step.</li>
</ol>
2. Preparation of Acute Brain Slice Cultures
2.1. Preparation of Solutions and Tools
<ol>
	<li>The following solutions are required (it is also possible to use high-glucose DMEM for dissecting and imaging17): 
	Dissection Medium (500 ml) 
	Gey&#39;s Balanced media - 500ml 
	45% Glucose - 5ml 
	Movie Medium (10 ml) 
	45% Glucose - 0.110 ml 
	HEPES&#160;1 M - 0.100 ml 
	Pen/Strep - 0.100 ml 
	FCS - 0.500 ml 
	B27 - 0.100 ml 
	Glutamine - 0.200 ml 
	Dulbecco Modified Eagle Medium (phenol red-free) - 8.89 ml</li>
	<li>Warm up movie medium at 37 &#176;C.</li>
	<li>Place Millicell inserts into a 35 mm glass bottom culture dish containing 1 ml of movie medium and place in a humidified incubator at 37 &#176;C/5% CO2.</li>
	<li>Prepare vibratome accessories (screwdriver, chamber, razor blades, glue).</li>
	<li>Prepare dissection tools: scissors, small spatula, straight forceps.</li>
	<li>Prepare tools for handling slices: small paintbrush or soft inoculating loop (size: 10 &#181;l), plastic Pasteur pipettes, an ice box and several 6 cm plastic dishes.</li>
	<li>Precool the dissection medium at 2-4 &#176;C.</li>
</ol>
2.2. Preparation of Brain Slices
<ol>
	<li>Fill a 6 cm dish with precooled dissection medium and place it on ice (to keep freshly cut slices).</li>
	<li>Following cervical dislocation, use scissors to decapitate the mouse pup. Remove the scalp with a scalpel, cut the skull along the mid-sagittal suture from the OB to the cerebellum and gently remove the cranial flaps using forceps. Make sure the entire brain is exposed and carefully dissect it out using a spatula, taking special care not to damage the tissue. The dissection has to be done carefully but at the same time very quickly (less than a minute if possible).&#160;Cervical dislocation is our preferred method because terminal anesthesia with drugs/gas anesthetics can influence the migratory properties of the neuroblasts and the healthy state of the brain slice cultures, which need to be imaged relatively quickly following animal sacrifice.</li>
	<li>Hemisect the brain with a razor blade (Figure 2). Discard uninjected hemisphere or use for other experiments.</li>
	<li>Place a small piece of tape on the vibratome holder and use the minimal necessary amount of glue to attach the brain hemisphere on top of it (Figure 2). This prevents damaging of the holder surface due to repeated glue applications.</li>
	<li>Let the glue dry for a few seconds.</li>
	<li>Place holder in the vibratome tray filled with precooled dissection solution. Point the olfactory bulb towards the blade (Figure 2).</li>
	<li>Begin cutting the brain hemisphere using appropriate settings. The following parameters are recommended: slice thickness 300 &#181;m; speed ~3-5; frequency ~9. High frequency and low speed are recommended to prevent damage to the slice.</li>
	<li>Collect slices using a small paintbrush or a soft inoculating loop. Only keep slices with visible OB (usually 2-3 slices/brain). Typically, the slice containing most of the RMS can be found at ~300 &#181;m from the bottom surface.</li>
	<li>Check slices under a standard fluorescent microscope for GFP signal (making sure to flip them over to check fluorescence on both sides), and choose the ones showing bright fluorescence along most of the RMS.</li>
</ol>
2.3. Culturing Brain Slices
<ol>
	<li>In a cell culture hood, cut away the most caudal third of the brain slice and remove any glue traces using fine straight tweezers or a microdissection scalpel.</li>
	<li>Delicately aspirate the slice using a plastic Pasteur pipette (cut the tip of the pipette to create a bigger opening, this will avoid damaging the slice) and place it on the center of a prewarmed Millicell insert.</li>
	<li>Make sure the side with the brightest fluorescent signal is placed in contact with the Millicell insert (for imaging with an inverted microscope).</li>
	<li>Remove excess dissection solution on top of the insert with a pipette.</li>
	<li>Leave slice cultures to settle in a 37 &#176;C/5% CO2 incubator for at least for 1 hr before imaging.</li>
</ol>
3. Time-lapse Imaging of Neuroblast Migration
<ol>
	<li>At least 2 hr before imaging, turn on the Perkin Elmer UltraViewVoX confocal spinning disk system, inverted Nikon Ti-E microscope, Hamamatsu C10600-10B (ORCA-R2) cooled digital CCD camera, and heating system (Solent Scientific) set at the constant temperature of 37 &#176;C.</li>
	<li>Open the Volocity software Acquisition module and click on &#34;VIz&#34; button and select the laser(s) for imaging.</li>
	<li>Within 2 hr of brain slice preparation, place the glass bottom dish containing the brain slice in the imaging chamber on the microscope stage.</li>
	<li>Use a Nikon CFI Super Plan Fluor ELWD 20X/0.45 objective under the appropriate fluorescent light to select and focus the area of the slice that will be imaged (e.g. the first descending part of the RMS just after the injection site, or the elbow region of the RMS, or just after the elbow before neuroblasts enter the OB).</li>
	<li>Set up the time-lapse imaging with the following actions:
	<ol>
		<li>On the microscope, click on the L100 button to allow laser scanning of the sample.</li>
		<li>In Volocity, open the UltraVIEW Laser Changer by selecting the appropriate laser (e.g. 488 nm laser for GFP).</li>
		<li>Set the exposure time (usually between 100-500 msec) and the laser intensity (usually 20-30%) depending on the intensity of the fluorescent signal.</li>
		<li>Adjust the image digital gain to improve cell visualization.</li>
		<li>Select the z-stack interval to image inside the brain slice (usually over a 100-120 &#181;m interval). Choose an interval with a suitable number of isolated cells to avoid possible overlaps as much as possible (this will facilitate the subsequent tracking analysis).</li>
		<li>Select the spacing between each z-stack image (usually 2-4 &#181;m).</li>
		<li>Choose the time interval between each z-stack capture (e.g. 3 min) and the total imaging duration (e.g. 3 hr).</li>
		<li>Click on the &#34;save&#34; icon to save the changes to the imaging parameters.</li>
		<li>Press the recording button to start imaging.</li>
	</ol>
	</li>
	<li>Alternate imaging of control and experimental samples (for example vehicle/drug treatment or different electroporated plasmids) throughout the same day.</li>
</ol>
4. Analyzing Neuroblast Migration
<ol>
	<li>In the Volocity Quantitation module, open the desired library created after completing a time-lapse experiment. Select &#34;Extended Focus&#34; from the top left box (Figure 4A, step 1).</li>
	<li>Click on the &#34;Measurements&#34; tab to display the Measurement window (Figure 4A, step 2).</li>
	<li>A list of tasks is visible in the bottom left of the screen. Drag &#34;Track&#34; (present under the &#34;Miscellaneous&#34; heading at the bottom of the list) on the space above. This will prompt the opening of a new window called &#34;Track&#34; at the top left section of the screen (Figure 4A, step 3).</li>
	<li>Select &#34;Points&#34; from the &#34;Input&#34; tab in the Track window (Figure 4A, step 4).</li>
	<li>Click on the Point Tool (Figure 4A, step 5).</li>
	<li>Start tracking the migrating neuroblast by clicking with the mouse on the central area of the cell body and keep tracking the cell movement until the last time point of the time-lapse is reached (for example point number 61 for a 3 hr-long movie).</li>
	<li>To obtain data choose &#34;Make Measurement Item&#34; from the Measurements Menu (Figure 4B, steps 6-7). A window will appear at the center of the screen.</li>
	<li>In this window, select &#34;A new measurement item called:&#34; and type a name (Figure 4C, step 8). Remember to select the &#34;All timepoints&#34; option before pressing OK (Figure 4C, step 9). A measurement item file will appear under the time-lapse file and will contain parameters for quantitative analysis (e.g. migration distance, velocity, displacement, and displacement rate).</li>
	<li>Double-click on the measurement item file to open it as a window (Figure 4D, step 10).</li>
	<li>To visualize the single tracks of analyzed cells, choose &#34;Tracked Point&#34; from the &#34;Display&#34; options (Figure 4D, step 11).</li>
	<li>Right click on the Measurement Item file and export it as a text file, which can then be imported in programs like Excel to analyze migration parameters.</li>
	<li>To measure movements between consecutive frames and pauses made by each cell during the filming period, in the Measurement Item file select from the &#34;Display&#34; options &#34;Point&#34; or &#34;Populations&#34; and click on ID (every track has an unique ID). Export the resulting file by proceeding as explained in step 4.10.</li>
</ol>

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T., McConnell, S. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytoskeletal+coordination+during+neuronal+migration.">Cytoskeletal coordination during neuronal migration.</a> Proc. Natl. Acad. Sci. U.S.A. 102, 13652-13657 (2005).</li><li>Valiente, M., Marin, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+migration+mechanisms+in+development+and+disease.">Neuronal migration mechanisms in development and disease.</a> Curr.Opin. Neurobiol. 20, 68-78 (2010).</li><li>Comte, I., 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=Galectin-3+maintains+cell+motility+from+the+subventricular+zone+to+the+olfactory+bulb.">Galectin-3 maintains cell motility from the subventricular zone to the olfactory bulb.</a> J. Cell Sci. 124, 2438-2447 (2011).</li><li>Sonego, 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=Fascin+regulates+the+migration+of+subventricular+zone-derived+neuroblasts+in+the+postnatal+brain.">Fascin regulates the migration of subventricular zone-derived neuroblasts in the postnatal brain.</a> J. Neurosci. 33, 12171-12185 (2013).</li></ol>

The authors have nothing to disclose.

Efficient migration of neural progenitors along the RMS ensures their subsequent maturation into functional neurons10. Prominent streams of neural progenitors directed towards the OB are visible in human infancy and are likely to play an important role in early postnatal human brain development27. Moreover, these cells are able to target sites of the brain affected by injury and neurodegeneration4,28. Being able to monitor in real time the effect of gene manipulation on neuroblast dynamics becomes essential to fully understand how neuroblast movement is guided and regulated.
Here we have described a protocol to monitor SVZ-derived neuroblast migration by coupling in vivo postnatal electroporation with time-lapse confocal spinning disk microscopy of acute brain slice cultures. Specifically, we have shown how migrating neural progenitors can be labeled by electroporating in the SVZ a plasmid encoding GFP. Depending on the purpose of the study, several types of plasmids can be used (e.g. allowing expression of other fluorescent proteins or co-expression of wild type/mutant proteins of interest, CRE recombinase, or shRNA together with fluorescent proteins). We strongly recommend using plasmids containing the chicken beta actin CAG promoter29 for in vivo expression. Imaging of brain slices obtained from electroporated animals can also be used to monitor the radial migration of neural progenitors in the OB at longer time points (7-10 days) after electroporation30.
After an initial period of practice, in vivo postnatal electroporation becomes a very reliable method, allowing robust neuroblast labeling for both time-lapse imaging and immunofluorescence analysis. Moreover, this technique offers a fundamental advantage over transgenic mice expressing fluorescent proteins under neuroblast-specific promoters, where the majority of neuroblasts are labeled. Indeed, electroporation allows sparse labeling of neuroblasts, thus allowing detailed analysis of their morphology and migration dynamics. Compared to the stereotactic delivery of viral vectors31, this technique is cheaper, faster and is very well tolerated by mouse pups. The main drawback consists in the fact that it is limited to early postnatal stages. Indeed, we strongly recommend using postnatal day 2 mouse pups, since we observed a substantial decrease in labeling efficiency at later times, when viral delivery methods become more suitable31. However, strategies like electroporation of CRE-expressing plasmids in appropriate mouse genetic models may be used to study neurogenesis in adult stages22.
Two-photon, standard confocal, spinning disk confocal, and wide-field fluorescence microscopy can all be used to visualize neuroblast migration17,23,24. Spinning disk confocal microscopy is a cheaper alternative to two-photon microscopy. It allows 3D imaging at higher speed through multiple z planes, limiting photobleaching compared to standard confocal microscopy and offering a higher resolution than wide-field fluorescence imaging31-33. The majority of migrating neuroblasts have a clearly visible soma and a highly dynamic leading process tipped with a growth cone.
As described by others17, having a perfusion chamber would allow to directly assess the effects of control and drug treatments on the same brain slice. While we consider this an important advantage, the protocol described here shows that it is not absolutely necessary for cell viability to perfuse slices with oxygenated artificial cerebrospinal fluid (aCSF) or high glucose DMEM17,33. Moreover, using an inverted microscope allows easier objective manipulation compared to upright microscopes equipped with water immersion objectives. We found that using a 20X long-distance objective is an optimal compromise, enabling tracking of a good number of cells (usually 25-40 per slice) and producing good resolution images (Video 1). Automatic tracking is also possible, however it is not always reliable. For this reason we recommend visual and, if necessary, manual verification of automatic tracking data. Higher magnification imaging of single neuroblasts can be performed using a Nikon Fluor DIC M/N2 40X/0.80W objective. The freely available ImageJ plugin MTrackJ can also be used to measure basic track statistics33, however some raw data acquisition files may be incompatible with this software and may need to be converted to suitable formats for analysis, which in certain cases can be time-consuming. The combination of image acquisition and analysis modules provided by the Volocity software allows an immediate transition from image capturing/processing to quantitative analysis of migratory parameters (speed, displacement, etc.), which can be readily visualized in a variety of graphs (e.g. showing migratory patterns/distance/directionality/speed/persistence/time spent immobile, etc.).
Neuroblasts display a saltatory movement, alternating migratory to stationary phases34,35 (Figure 3C). In light of the fact that stationary phases can be between 4-10 min32, we believe that capturing images every 3 min represents a good compromise to follow migrating neuroblasts with minimum illumination and photodamage. We rarely see cells stopping for more than 6 min, and find that, over a 3-hr period, they generally spend an average of 30 min immobile. According to previous reports, neuroblasts have an average speed of 60-100 &#181;m/hr and an average meandering index (actual displacement over the migrated distance), slightly above 0.623,36, which is in agreement with our observations. Typically, about 60% of cells show a &#34;migratory&#34; behavior (e.g. following almost linear migration paths, with a meandering index between 0.6-1) and 20-25% are &#34;exploratory&#34; (with a meandering index between 0-0.4), while the remaining ~20% can be classified as &#34;intermediate&#34; (alternating migratory and exploratory phases, with a meandering index between 0.4-0.6).
We recognize that there are still limitations to this technique, since we cannot film neuroblasts for more than 3-4 hr without observing substantial alterations in their motility. Filming duration could be increased by lowering imaging frequency (however this will affect the accuracy of the migration analysis) as well as by adopting a perfusion chamber to optimize environmental conditions for imaging. However, the protocol described here including a 3-4 hr imaging period represents a suitable compromise enabling the collection of sufficient data for quantitative analysis of migration. Indeed, so far we have managed to detect the effects produced on migration by protein overexpression/downregulation37 or by pharmacological manipulation after comparison with appropriate controls (Sonego and Zhou, unpublished results).
In conclusion, the combination of in vivo postnatal electroporation with spinning disk imaging of brain slice cultures represents a powerful tool to monitor the dynamics of neuroblast migration in a system closely resembling the in vivo environment, offering the possibility to investigate in depth the molecular mechanisms controlling neuroblast motility.

In the mammalian brain, the generation of new neurons (neurogenesis) occurs after birth mainly in two regions, the subventricular zone (SVZ) of the lateral ventricles and the subgranular zone in the dentate gyrus of the hippocampus1. Considerable evidence gathered in recent years supports a critical role for postnatal neurogenesis in hippocampal and olfactory bulb memory functions1-3. Importantly, postnatal neurogenesis also holds therapeutic potential because of its relationship with degenerative neurological disorders, and the ability of neuroblasts to migrate to injured sites in the brain4-6.
The subventricular zone (SVZ) has recently emerged as a crucial neurogenic niche. SVZ-derived neuroblasts migrate towards the olfactory bulb (OB) via the rostral migratory stream (RMS), making this the longest migration process in the postnatal brain1,7,8. The mammalian SVZ/RMS/OB system has become a useful model to study different steps in neurogenesis, such as proliferation, migration and differentiation1,8. Many growth factors and extracellular cues regulate SVZ neurogenesis and migration along the RMS, but the intracellular molecular mechanisms are far from being fully understood1,9. Proper migration along the RMS is crucial for the subsequent maturation of newborn neurons10. Additionally, some studies have shown that SVZ-derived neuroblasts can migrate out of the RMS to brain injury sites4-6,11-13. Thus, investigating the signalling mechanisms regulating neuroblast migration is fundamental not only to understand neurogenesis but also for potential therapeutic applications.
Here, we describe a detailed protocol to label SVZ neural progenitors by in vivo postnatal electroporation and monitor their migration along the RMS in acute brain slice cultures using time-lapse spinning disk confocal microscopy. Electroporation is widely used in developmental studies from embryonic to adult stages14-18. It is a powerful tool to target and manipulate SVZ neural progenitors and represents a cheaper and considerably faster alternative to stereotactic injection of viral vectors or generation of transgenic models1,15,19,20. It is a relatively simple procedure that does not need surgery and has high survival rates. Electroporation of shRNA or CRE recombinase-expressing plasmids in mouse genetic models employing the LoxP system can be used to target genes of interest or to achieve permanent labeling of SVZ progenitors, thus representing a useful tool for adult neurogenesis studies21,22.
Imaging RMS neuroblast migration in the intact brain is still challenging due to current technical limitations. However, this process can be monitored using confocal spinning disk time-lapse microscopy of acute brain slices, which provide a suitable system closely resembling the in vivo condition also amenable to pharmacological manipulation23,24. Coupling in vivo postnatal electroporation with time-lapse imaging will facilitate the understanding of the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

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			<td height="21" style="height:21px;width:344px;">Millicell</td>
			<td style="width:195px;">Millipore</td>
			<td style="width:149px;">PICM0RG50</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">35 mm Glass bottom culture dish</td>
			<td>MatTek</td>
			<td style="width:149px;">P35G-0-14-C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gey&#39;s Balanced media</td>
			<td style="width:195px;">Sigma</td>
			<td>G9779-500ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glucose, 45%</td>
			<td style="width:195px;">Sigma</td>
			<td>G8769-100ML</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HEPES</td>
			<td style="width:195px;">Sigma</td>
			<td>H3375-25G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pen/Strep</td>
			<td style="width:195px;">GIBCO</td>
			<td style="width:149px;">15140-122</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">FCS</td>
			<td style="width:195px;">GIBCO</td>
			<td style="width:149px;">10109-163</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">B27 supplement</td>
			<td style="width:195px;"><a>Invitrogen Life Technologies</a></td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">L-Glutamine</td>
			<td style="width:195px;"><a>Invitrogen Life Technologies</a></td>
			<td>25030-081</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DMEM (phenol red-free)</td>
			<td style="width:195px;">GIBCO</td>
			<td style="width:149px;">31053-028</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fast Green</td>
			<td style="width:195px;">Sigma</td>
			<td>F7252-5G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Glass capillaries for injection</td>
			<td style="width:195px;">Harvard Apparatus</td>
			<td style="width:149px;">30-0057</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Aspirator tube</td>
			<td style="width:195px;">Sigma</td>
			<td style="width:149px;">A5177</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sutter P-97 capillary puller</td>
			<td style="width:195px;">Sutter Instrument</td>
			<td style="width:149px;">P-97</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">ECM830 Square Wave Electroporator</td>
			<td style="width:195px;">Harvard Apparatus</td>
			<td style="width:149px;">45-0052</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Platinum Tweezertrodes 7 mm</td>
			<td style="width:195px;">Harvard Apparatus</td>
			<td style="width:149px;">45-0488</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Footswitch Model 1250F</td>
			<td style="width:195px;">Harvard Apparatus</td>
			<td style="width:149px;">45-0211</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Gel for electrodes</td>
			<td style="width:195px;">Cefar Compex</td>
			<td style="width:149px;">6602048</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isoflurane</td>
			<td style="width:195px;">Merial</td>
			<td style="width:149px;">AP/DRUGS/220/96</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vibratome</td>
			<td style="width:195px;">Leica</td>
			<td style="width:149px;">VT1000S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glue</td>
			<td>Roti coll</td>
			<td>Roti coll 1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">UltraViEW VoX spinning disk system</td>
			<td style="width:195px;">Perkin Elmer</td>
			<td style="width:149px;"></td>
			<td>Customized setup (multiple laser sources can be used) equipped with Hamamatsu ORCA R2 C10600-10B CCD camera</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Volocity software</td>
			<td style="width:195px;">Perkin Elmer</td>
			<td style="width:149px;"></td>
			<td>Acquisition, Quantitation, Visualization Modules</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Environmental chamber for microscopy</td>
			<td style="width:195px;">Solent Scientific</td>
			<td style="width:149px;"></td>
			<td>Custom-made</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:344px;">Ti-E inverted microscope</td>
			<td style="width:195px;">Nikon</td>
			<td style="width:149px;"></td>
			<td>CFI Super Plan Fluor ELWD 20X/0.45 NA objective is recommended for the application described in this paper</td>
		</tr>
	</tbody>
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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.

Labeling of SVZ-derived migratory neuroblasts can be observed along the RMS, usually 4-8 days after a successful electroporation (Figure 1B). Longer time points can also be chosen, but less cells will be found in the RMS since most of them will have entered the OB. Neuroblasts start acquiring typical morphology and features of mature granule cells in the OB around 2-3 weeks after electroporation (not shown). After culturing for ~1 hr, brain slices from electroporated mouse pups can be reliably imaged for up to 3-4 hr. As previously reported23,26, neuroblasts can display complex migration dynamics (Figure 3 and Video 1), which can be quantitatively analyzed using cell tracking (Figure 4).
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50905/50905fig1.jpg" width="500px" /> 
Figure 1. Postnatal in vivo electroporation. (A) Schematic drawing of in vivo electroporation of a postnatal day 2 mouse pup. A dotted line (red) connecting the eye to the craniometrical landmark lambda serves as a positional marker for capillary insertion. The injection point is indicated as a green dot. Grey oval shapes indicate electrode position as described in Boutin et al.15&#160;(B) Sagittal mouse forebrain slice immunostained for GFP 5 days after electroporation of a GFP-expressing plasmid. SVZ-derived migrating neuroblasts are visible along the RMS and some of them have started to reach the OB. Ctx: cortex; SVZ: subventricular zone; RMS: rostral migratory stream; OB: olfactory bulb. Bar, 400 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50905/50905fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/50905/50905fig2.jpg" width="500px" /> 
 
Figure 2. Schematic steps in the preparation of acute brain slice cultures for imaging. (A) Caudal and intrahemispheric cuts (dotted lines) are performed on a freshly dissected brain. (B) The electroporated brain hemisphere is placed onto a vibratome holder. (C) Sagittal slices are obtained with a vibratome and observed under a standard fluorescent microscope. (D) Slices with the best GFP signal are cultured for at least 1 hr on a Millicell insert inside a glass-bottom dish, and (E) subsequently placed in an environmental chamber for imaging by an inverted confocal spinning disk system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50905/50905fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50905/50905fig3.jpg" width="500px" /> 
Figure 3. Time-lapse imaging of migrating neuroblasts. (A) Spinning disk time-lapse images of neuroblasts taken from a mouse sagittal brain slice 5 days after electroporation of a GFP-expressing plasmid. Images are 1 hr apart. Each panel is a z-stack projection of 28 consecutive images 4 &#181;m apart. Arrowheads indicate 4 representative neuroblasts migrating along the RMS towards the olfactory bulb (located out of the picture at the bottom right corner). (B) Representative migratory paths obtained from time-lapse imaging of the 4 neuroblasts highlighted in (A). (C) Graph showing the distance migrated with time by 2 representative neuroblasts. Cells display a typical saltatory motile behavior. Bar, 70 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50905/50905fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="3in" src="/files/ftp_upload/50905/50905fig4.jpg" width="400px" /> 
Figure 4. Tracking analysis of migrating neuroblasts. Sequential steps used to track migrating neuroblasts using Volocity software. Please see text for detailed description. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/50905/50905fig4highres.jpg" target="_blank">Click here to view larger image</a>.
Video 1: Time-lapse imaging of migrating neuroblasts. The movie shows a section of the mouse RMS with GFP-labeled migrating neuroblasts obtained 5 days after electroporation of a GFP-expressing plasmid. The OB is located out of view towards the bottom right corner. Confocal z-stacks were captured on a spinning disk confocal with a 20X objective every 3 min for 3 hr over a 120 &#181;m interval. Playing speed: 10 frames/sec.

Watch this Scientific Journal Video about In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices at JoVE.com

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.

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 ...

in-vivo-postnatal-electroporation-time-lapse-imaging-neuroblast

Nanyang Technological University

Research

JoVE Journal

Neuroscience

13.1K Views.  King's College London. 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.

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

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 ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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 ...

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then radial migration in the molecular layer and the Purkinje cell layer to reach the internal granular layer of the cerebellar cortex. Default in migratory processes induces either cell death or misplacement of the neurons, leading to deficits in diverse cerebellar functions. Centripetal granule cell migration involves several mechanisms, such as chemotaxis and extracellular matrix degradation, to guide the cells towards their final position, but the factors that regulate cell migration in each cortical layer are only partially known. In our method, acute cerebellar slices are prepared from P10 rats, granule cells are labeled with a fluorescent cytoplasmic marker and tissues are cultured on membrane inserts from 4 to 10 hr before starting real-time monitoring of cell migration by confocal macroscopy at 37 &#176;C in the presence of CO2. During their migration in the different cortical layers of the cerebellum, granule cells can be exposed to neuropeptide agonists or antagonists, protease inhibitors, blockers of intracellular effectors or even toxic substances such as alcohol or methylmercury to investigate their possible role in the regulation of neuronal migration.

Ex Vivo Imaging of Postnatal Cerebellar Granule Cell Migration Using Confocal Macroscopy

During postnatal cerebellum development, immature granule cells originating from the germinal zone exhibit distinct modalities of migration to reach their final destination and to establish neuronal networks. This protocol describes the preparation of cerebellar slices and the confocal macroscopic approach used to investigate the factors that regulate neuronal migration.

During postnatal development, immature granule cells (excitatory interneurons) exhibit tangential migration in the external granular layer, and then ...

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It has helped to describe the different steps of radial migration and characterize the molecular mechanisms controlling this process. To directly and dynamically analyze migrating neurons they have to be traced over time. This protocol describes a workflow that combines in utero electroporation with organotypic slice culture and time-lapse confocal imaging, which allows for a direct examination and dynamic analysis of radially migrating cortical neurons. Furthermore, detailed characterization of migrating neurons, such as migration speed, speed profiles, as well as radial orientation changes, is possible. The method can easily be adapted to perform functional analyses of genes of interest in radially migrating cortical neurons by loss and gain of function as well as rescue experiments. Time-lapse imaging of migrating neurons is a state-of-the-art technique that once established is a potent tool to study the development of the cerebral cortex in mouse models of neuronal migration disorders.

Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation

This protocol provides instructions for direct observation of radially migrating cortical neurons. In utero electroporation, organotypic slice culture, and time-lapse confocal imaging are combined to directly and dynamically study the effects of overexpression or downregulation of genes of interest in migrating neurons and to analyze their differentiation during development.

In utero electroporation is a rapid and powerful approach to study the process of radial migration in the cerebral cortex of developing mouse embryos. It ...