We thank Jacqueline Andratschke, Elena Werle, Sachi Takenaka, and Matthias Toberer for excellent technical assistance, as well as Victor Tarabykin for helpful discussions. This work was supported by a grant of the Deutsche Forschungsgemeinschaft to S.B. (BR-2215).

All experimental procedures were approved by the Animal Welfare Committee (Regierungspr&#228;sidium T&#252;bingen) and carried out in accordance by the German Animal Welfare Act and the EU Directive 2010/63/EU.
1. In Utero Electroporation
<ol>
	<li>Microinjection needles
	<ol>
		<li>Pull borosilicate glass capillaries (outer diameter: 1.0 mm, inner diameter: 0.58 mm, length: 100 mm) into microinjection needles using a micropipette puller with a box filament (2.5 mm x 2.5 mm) and the following program: HEAT: 540, PULL: 125, VELOCITY: 20, and DELAY: 140. Determine the HEAT value for every individual filament by performing a RAMP test (HEAT value: RAMP+25).</li>
		<li>Bevel needles at an angle of 38&#176; and intermediate to high revolution speed with a microgrinder to obtain a tip size of 20 to 30 &#181;m for injecting at embryonic day (E) 13.5 or younger and 30 to 40 &#181;m for injecting at older embryonic stages. For storage, fix needles in a box or petri dish with modeling clay to prevent damage to the tips.</li>
	</ol>
	</li>
	<li>Plasmid DNA Solution
	<ol>
		<li>Prepare plasmid DNA construct containing fluorescent reporter protein (e.g. GFP) using an endotoxin-free maxi preparation kit according to the manufacturer&#39;s instructions.</li>
		<li>Dilute plasmid DNA solution to a final concentration of 1 - 2 &#181;g/&#181;L in endotoxin-free Tris-EDTA buffer containing Fast Green at a final concentration of 0.01%. Aliquot solution and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Bacteriostatic Sodium Chloride Solution
	<ol>
		<li>Prepare bacteriostatic sodium chloride solution by adding benzyl alcohol to a final concentration of 0.9% to isotonic 0.9% sodium chloride solution. Sterile filter solution immediately before use.</li>
	</ol>
	</li>
	<li>Carprofen Injection Solution
	<ol>
		<li>Prepare carprofen injection solution by adding carprofen to a final concentration of 0.5 mg/mL to sterile isotonic 0.9% sodium chloride solution.&#160;Store at 4 &#176;C for up to 28 days.</li>
	</ol>
	</li>
	<li>Anesthesia, DNA Injection, and Electroporation
	<ol>
		<li>Weigh an E14.5 timed-pregnant mouse and place it into a transparent anesthetizing chamber connected to a vaporizer and saturated with 5% isoflurane. When the animal is unconscious, transfer it to an anesthetizing mask attached to a 37 &#176;C warming plate and keep it anesthetized with 1.8 - 2.2% isoflurane. 
		Note: Assess the development of surgical anesthesia by loss of tail pinch and pedal reflexes (toe pinch).</li>
		<li>For analgesia, inject 100 &#181;L of carprofen injection solution per 10 mg of mouse bodyweight (5 mg/kg bodyweight) subcutaneously. Turn the mouse with its back down and carefully cover the eyes with petroleum jelly to prevent them from drying.</li>
		<li>Gently spread out the limbs and fix them to the warming plate with surgical tape. Sterilize the abdomen with 70% ethanol followed by iodine solution (7.5 mg/mL) using cellulose swabs.&#160;Repeat this procedure three times to ensure for proper disinfection.</li>
		<li>Cover the abdomen with sterile gauze in which an incision has been cut to spare out a field (diameter: 2 - 2.5 cm) for the abdominal incision. Moisten the gauze with bacteriostatic sodium chloride solution. 
		Caution: Reassess the development of surgical anesthesia by loss of the pedal reflex.</li>
		<li>Using Micro Adson Forceps (serrated, length: 12 cm) and fine scissors (angled to side, length: 9 cm) cut the skin approximately 1.5 cm along the midline of the abdomen. Cut the underlying muscle along the linea alba.</li>
		<li>Remove one uterine horn with ring forceps (length: 9 cm, outer diameter: 3 mm, inner diameter: 2.2 mm) and gently place it onto the moistened gauze. 
		Caution: Hold the uterine horn with ring forceps only in between embryos and do not perturb the placentas or any supply vessels. Keep the uterus moist at all times with bacteriostatic sodium chloride solution.</li>
		<li>Carefully position an embryo with ring forceps and locate the lateral ventricle, which presents as a crescent shaped shadow parallel to the sagittal sinus. The injection site lies approximately in the middle of a line between the pigmented eye and the confluence of sinuses, where the sagittal sinus branches to two transverse sinuses. Push the microinjection needle through the uterine wall and into the lateral ventricle. 
		Note: If necessary, the depth of the injection can be corrected by retraction of the needle.</li>
		<li>Inject 1 to 2 &#181;L of DNA solution into the lateral ventricle with a microinjector, which is operated with a foot pedal, using the following settings: 25 psi, 10 to 20 ms per pulse, and 5 to 10 pulses. Successful injection can be monitored by the colored DNA solution, which should fill the ventricular system.</li>
		<li>Place tweezers-type electrodes (3 mm diameter for E13.5 or younger and 5 mm diameter for E14.5 or older) in such a way that the &#39;positive&#39; terminal is on the same side as the injected ventricle and the &#39;negative&#39; terminal is on the opposite side of the injected ventricle below the ear of the embryo&#39;s head.</li>
		<li>Moisten the electroporation site with a few drops of bacteriostatic sodium chloride solution and apply 5 electrical current pulses (35 V for E13.5 or younger and 40 V for E14.5 or older) of 50 ms duration with 950 ms intervals. 
		Note: Currents of 60 to 90 mA are usually sufficient for successful electroporation. Lower currents will not efficiently transfect neurons, whereas higher currents may lead to embryonic death.</li>
		<li>Repeat the procedure (c.f. Step 1.5.7. to 1.5.10.) for every embryo of the uterine horn and gently place it back into the abdominal cavity.</li>
		<li>Remove the uterine horn from the other side and repeat the procedure described in Step 1.5.7. to 1.5.11.</li>
		<li>Close the abdominal cavity by suturing the muscle layer before suturing the skin using Micro Adson Forceps, Mathieu Needle Holder (tungsten carbide, length: 14 cm), and non-absorbable surgical suture (3/8 circle, 13 mm, taper point). 
		Caution: Be careful not to capture the uterus or any other organs during suture.</li>
		<li>Carefully disinfect the skin suture with iodine solution (7.5 mg/mL) and gently remove periocular excess of petroleum jelly using cellulose swabs.&#160;&#160;Place the mouse under an infrared lamp until it wakes up. Closely monitor the mouse during the next two days. Repeat the analgesia treatment as described in Step 1.5.2 after 12 to 24 h.</li>
	</ol>
	</li>
</ol>
2. Organotypic Slice Culture
<ol>
	<li>Laminin Stock Solution
	<ol>
		<li>Dissolve 1 mg of lyophilized laminin in sterile water to obtain a 1 mg/mL laminin stock solution. Prepare 50 &#181;L aliquots and store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Poly-L-Lysine Stock Solution
	<ol>
		<li>Dissolve 50 mg of poly-L-lysine in sterile water to obtain a 1 mg/mL poly-L-lysine stock solution. Prepare 0.5 mL aliquots and store at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Complete Hank&#39;s Balanced Salt Solution (Complete HBSS)
	<ol>
		<li>Prepare complete HBSS by combining 50 mL of 10x HBSS solution, 1.25 mL of 1 M HEPES buffer (pH 7.4), 15 mL of 1 M D-glucose solution, 5 mL of 100 mM calcium chloride solution, 5 mL of 100 mM magnesium sulfate solution, and 2 mL of 1 M sodium hydrogen carbonate solution. Add sterile water up to 0.5 L, sterile filter, and store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Slice Culture Medium
	<ol>
		<li>Combine 35 mL of Basal Medium Eagle (BME), 12.9 mL complete HBSS (c.f. section 2.3.1.), 1.35 mL of 1 M D-glucose, 0.25 mL of 200 mM L-glutamine, and 0.5 mL of penicillin-streptomycin to obtain 50 mL of slice culture medium. Sterile filter and add horse serum to a final concentration of 5%. Store at 4 &#176;C for up to 4 weeks.</li>
	</ol>
	</li>
	<li>Low Melting Point (LMP) Agarose Solution
	<ol>
		<li>Prepare 3% LMP agarose solution by dissolving 1.5 g of LMP agarose in 50 mL of complete HBSS by heating in a microwave oven for 1 to 2 min at intermediate power. 
		Note: Monitor heat carefully to prevent overflow during boiling.</li>
		<li>Place solution in a water bath at 38 &#176;C until ready for use. Store at 4 &#176;C and reuse once.</li>
	</ol>
	</li>
	<li>Coating of Membrane Inserts
	<ol>
		<li>Add one aliquot of laminin stock solution (c.f. Step 2.1.1.) and one aliquot of poly-L-lysine stock solution (c.f. Step 2.2.1.) to 6 mL of sterile water to obtain coating solution (enough for 6 inserts).</li>
		<li>Place membrane inserts into a 6-well plate containing 2 mL of sterile water in the bottom of each well. Add 1 mL of coating solution (c.f. Step 2.6.1.) on top of each membrane and incubate overnight at 37 &#176;C and 5% carbon dioxide atmosphere. 
		Note: Only use inserts with optically clear membranes, such as polytetrafluoroethylene (PTFE) membrane.</li>
		<li>Wash the membranes three times with 1 mL of sterile water and dry. Use coated membranes on the same day or store in a clean 6-well plate at 4 &#176;C for up to four weeks.</li>
	</ol>
	</li>
	<li>Brain Dissection and Embedding
	<ol>
		<li>Two days after the electroporation place the mouse into a transparent chamber connected to a vaporizer and saturated with 5% isoflurane. When the animal is unconscious, remove it from the chamber and sacrifice it by cervical dislocation.</li>
		<li>Remove the uterus containing the embryos and place it into a 10-cm Petri dish with ice-cold complete HBSS. 
		Note: From this point onward keep all solutions, embryos, and brains on ice.</li>
		<li>Use a stereomicroscope, fine tipped forceps (straight, 11 cm) and a pair of Vannas T&#252;bingen Spring Scissors (angled up, length: 9.5 cm) to separate each embryo from the uterus and transfer it to another petri dish containing ice-cold complete HBSS.</li>
		<li>Make an incision at the level of the brain stem and cut along the sagittal midline. Peel away the skin and cartilage covering the brain. Then, cut off the brain stem right behind the hemispheres and remove the brain from the skull. Transfer the brain with a micro-spoon spatula (185 mm x 5 mm) to a 12-well plate containing ice-cold complete HBSS. Collect all brains from the litter in the 12-well plate.</li>
		<li>Use a fluorescent stereomicroscope to screen the brains in the 12-well plate for fluorescence brightness as well as size of the electroporated region. Select 2 to 4 brains with bright fluorescent signals in the presumptive somatosensory cortex for further processing.</li>
		<li>Pour 3% LMP agarose solution kept at 38 &#176;C into peel-away disposable embedding mold. Remove brain from the 12-well plate with a micro-spoon spatula and carefully drain excess complete HBSS around the brain using fine tissue paper.</li>
		<li>Place the brain gently into the agarose solution, push it to the bottom of the mold, and carefully adjust its position using a 20-gauge needle. For coronal sections, orient the brain with the olfactory bulbs pointing upwards. Keep the mold on ice until the agarose solution is solidified and continue with sectioning the brain.</li>
	</ol>
	</li>
	<li>Vibratome Sectioning and Slice Culture
	<ol>
		<li>Remove the agarose block from the mold and place it onto the lid of a sterile petri dish. Trim away excess agarose with a clean razor blade leaving approximately 2 to 3 mm below the olfactory bulbs and 1 mm of agarose on the other sides. Fix the trimmed agarose block with the olfactory bulbs pointing downwards to a specimen stage using&#160;cyanoacrylate glue. 
		Note: Sterilize instruments and equipment with 70% ethanol before sectioning.</li>
		<li>Transfer the specimen stage to the slicing chamber of a vibrating blade microtome containing ice-cold complete HBSS and position the brain with its dorsal side towards the blade. Cut 250 &#181;m thick brain slices containing the electroporated region with a low to moderate amplitude of 0.9 mm and a very slow sectioning speed of 0.09-0.12 mm/s. Usually 4 to 6 slices with bright fluorescent signal are obtained from each successfully electroporated brain.</li>
		<li>With a bent micro-spoon spatula and fine tipped forceps, carefully transfer the brain slices with the surrounding agarose to a 6-well plate containing ice-cold complete HBSS. 
		Caution: Be careful not to damage the brain tissue by only touching the agarose rim around the sections with the forceps during transfer.</li>
		<li>Process all selected brains as described in Steps 2.8.1. to 2.8.3.</li>
		<li>Moisten the membrane inserts with 100 &#181;L of complete HBSS before placing the brain slices onto the membranes to facilitate positioning of the slices. Up to 5 slices can be placed on 1 membrane insert.</li>
		<li>Carefully transfer the slices onto the membranes using a bent micro-spoon spatula and fine tipped forceps. Gently pull a slice onto the spatula by only touching the agarose rim with the forceps and gently push the slice from the spatula onto the membrane. Position the slice using forceps and continue to transfer the remaining slices. Remove excess complete HBSS with a pipette. 
		Note: Do not overlap the slices with one another.</li>
		<li>With the help of forceps carefully place the membrane inserts with the slices into a 6-well plate containing 1.8 mL of slice culture medium (c.f. Step 2.4.1.) and incubate at 37 &#176;C and 5% carbon dioxide atmosphere until time-lapse imaging.</li>
	</ol>
	</li>
</ol>
3. Time-Lapse Imaging
<ol>
	<li>Microscope Setup
	<ol>
		<li>Set a climate chamber placed onto the stage of an inverted confocal microscope to 37 &#176;C and 5% carbon dioxide atmosphere. Use a hybrid detector to reduce laser intensity and improve cell viability. For detailed analysis, use an extra-long working distance 40X dry objective with a numerical aperture of 0.6 or higher. 
		Note: All components of the microscope need to be pre-warmed to 37 &#176;C for 2 - 3 h before imaging to provide a stable focus during the imaging process.</li>
	</ol>
	</li>
	<li>Slice Imaging
	<ol>
		<li>Select a brain slice for imaging from the 6-well plate containing the membrane inserts using an inverted fluorescence microscope. 
		Caution: Avoid prolonged exposure time to ultraviolet light as this may cause cellular photodamage. 
		Note: Select a slice with bright single neurons in the upper SVZ migrating radially towards the pial surface of the slice. Fine fluorescent processes of radial glial cells that span the entire CP indicate an intact radial glial scaffold, which is used by radially migrating cortical neurons.</li>
		<li>Transfer the membrane insert with the slice, which was selected for time-lapse imaging, into a 50 mm diameter glass bottom dish containing 2 ml of slice culture medium with the help of forceps. Place the dish into the climate chamber of the confocal microscope.</li>
		<li>Set the resolution to 512 by 512 pixels and increase the scan speed from 400 Hz to 700 Hz in order to increase the frame rate from about 1.4 to about 2.5 frames per second, respectively. Use no more than two times averaging. Define a z-stack through the electroporated region (usually 400-100 &#956;m) with a step size of 1.5 &#956;m. Collectively, these settings allow for a sufficient resolution and brightness of the image while keeping photodamage low during acquisition. Start the time-lapse series by taking a z-stack every 30 min. 
		Note: Prolonged exposure of the slice to laser light will induce photodamage reducing the viability of the cells. Adjust the exposure time to laser light accordingly.</li>
		<li>Analyze time-lapse series using ImageJ software (https://imagej.nih.gov/iij/) or equivalent.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Radial migration is a key process in neocortex development. Mutations in genes affecting different steps of this process can cause severe cortical malformations, including lissencephaly and white matter heterotopia1,2. We recently showed that Bcl11a, which is expressed in young migrating cortical projection neurons, plays a role in radial migration. We used time-lapse confocal imaging of migrating neurons in acute cortical slices of electroporated brains to directly demonstrate that genetic deletion of Bcl11a in migrating neurons causes polarization and migration defects. Time-lapse series were used to trace migration paths of individual neurons and calculate specific parameters of migrating neurons, including speed profiles, average migration speed, and migratory direction10.
This protocol describes an important approach when studying molecular mechanisms in radial migration. By using short hairpin RNA or DNA expression plasmids, almost any gene of interest can be knocked down or overexpressed, respectively. It is a powerful approach to combine electroporation with the Cre-LoxP system. Transfection of brains of conditional mutant (&#34;floxed&#34;) mice with Cre recombinase generates a mosaic mutant situation and allows selective analysis of single mutant neurons that are surrounded by wild-type cells. In this way, cell autonomous molecular mechanisms in migrating neurons can be investigated. Also, functional rescue experiments are easily performed. Preparing time-lapse series of electroporated brain slices allows dynamically analyzing migratory behavior of single migrating neurons, which provides much more information than can be obtained from fixed brain sections. In utero electroporation3,4 requires initial training but &#8211; once mastered &#8211; is a rapid and straightforward technique to reproducibly transfect neurons of the cerebral cortex in vivo. In the protocol described here we used in utero electroporation at E14.5, when late-born upper-layer neocortical projection neurons are born. For the study of early-born deep-layer projection neurons in utero electroporation has to be performed on earlier developmental stage (i.e. E12.5)13. In principle, the protocol may also be used to study migration of other neuronal subpopulations in the brain that can be transfected with electroporation, including migration of interneurons14 and cerebellar granule cells15.
The slice culture protocol that we describe was adapted from Polleux and Gosh12 and previously, we have used it for ex utero electroporation16,17 to study hippocampal development. Most importantly, the brain slices have to be handled with care throughout the procedure to maintain their viability. We use an inverted microscope and optical quality glass bottom dishes to image the slice culture resting on a cell culture insert. This configuration is very easy to set up and allows sterile conditions, since the objective never gets in touch with the slice culture or medium. However, long working distance objectives (&#62;2 mm) with a sufficiently high numerical aperture (0.6 or higher) are required. Other studies have put the brain slice directly onto the glass bottom and used a collagen matrix18,19 to fix the slice in place, which makes the use of objectives with a shorter free working distance possible. However, using cell culture inserts provides easy handling, reproducible results, and allows the researcher to image the slices even 1 to 2 days after their preparation without compromising viability of the brain slice (data not shown). Another critical issue concerns damaging the slices by laser light. We have used a highly sensitive hybrid detector, which allowed us to reduce laser power to a minimum while still obtaining sufficient fluorescent signals for time-lapse imaging. Using multiphoton imaging would further reduce photodamage and would allow for a deeper tissue penetration as well as more efficient light detection as compared to conventional confocal microscopy.
Despite its usefulness to study neuronal migration, the protocol has limitations and cannot entirely preserve the situation of migrating neurons in the intact brain. Extracellular matrix, the adhesive intercellular contacts, and the vasculature within the migratory zone can dynamically influence radially migrating neurons20,21. Specifically, upper layer neurons depend on elongated radial glial processes for their migration22. For successful imaging, it is therefore important to select brain slices with GFP electroporated radial glial cells that extend their processes through the entire cortical width. &#39;Oblique slices&#39;, in which the radial glial scaffold has been cut off before reaching the pial surface of the brain, should be discarded from further analysis. Sometimes, cell death may occur at the air-surface of the brain slice, which restricts image acquisition to deeper region within the sample. Electroporated neurons in cultured brain slices may migrate less efficiently compared to electroporated neurons in vivo, which can be caused by insufficient recovery from preparation. This requires analyses of multiple slice cultures for both experimental and control situations, and critical interpretation of representative data sets. In some cases the insults associated with in utero electroporation may cause embryonic death or structural alterations of the brain, including enlarged ventricles, asymmetrically thinned cortex, or a glial scar formation resulting from DNA injection. In those cases, the sample should be discarded from further analysis.
While brain slices from embryonic donors survive well on membrane inserts23, analyses are restricted to early developmental events, including neuronal migration, and we have not used it for studying later events, such as dendrite development or synaptogenesis. In summary, we provide a detailed protocol to directly and dynamically study neuronal migration of electroporated neurons in slice cultures, which can be readily adapted to analyze functions of genes involved in neurodevelopmental disorders.

The neocortex is the major site of cognitive, emotional, and sensorimotor functions. It is composed of six horizontal layers oriented in parallel to the surface of the brain. During development progenitor cells in the lateral wall of the dorsal telencephalon give rise to projection neurons that migrate radially towards the pial surface and acquire a layer type-specific neuronal identity. After being generated in the ventricular/subventricular zones (VZ/SVZ) these neurons become transiently multipolar and slow their migration. After a short stay in the intermediate zone (IZ) they switch to a bipolar morphology, attach to the radial glial scaffold, and continue radially oriented migration into the cortical plate (CP). Upon reaching their final target projection neurons detach from the radial glial processes and acquire layer-specific identity. Mutations in genes affecting different steps of neuronal migration can cause severe cortical malformation, such as lissencephaly or white matter heterotopia1,2.
In utero electroporation is a rapid and powerful technique to transfect neural progenitor cells in the developing brain of rodent embryos3,4. With this technique it is possible to knockdown and/or overexpress genes of interest in order to study their functions in developing neurons. This method has specifically helped to describe the morphological details and characterize the molecular mechanisms of the process of radial migration5,6,7,8,9. Radially migrating neurons undergo dynamic changes in cell shape, migration speed, as well as migratory direction, which require direct and continuous observation over time. Organotypic slice culture and time-lapse confocal imaging of electroporated brains allow to directly observe migrating neurons over time. Using this combined approach, it is possible to analyze distinct features of migrating neurons that cannot be investigated in fixed tissue sections of electroporated brains.
We recently applied time-lapse confocal imaging of migrating neurons in slice cultures of electroporated brains to study the role of the transcription factor B cell CLL/lymphoma 11a (Bcl11a) during cortical development10. Bcl11a is expressed in young migrating cortical neurons and we used a conditional mutant Bcl11a allele (Bcl11aflox)11 to study its functions. Electroporation of Cre recombinase together with green fluorescent protein (GFP) into cortical progenitors of Bcl11aflox/flox brains allowed us to create a mosaic mutant situation, in which only few cells are mutated in an otherwise wild-type background. In this way, it was possible to study cell-autonomous functions of Bcl11a at the single cell level. We found that Bcl11a mutant neurons display reduced speed, shifts in their speed profiles, as well as random-like orientation changes during their migration10. In the outlined protocol we describe a workflow for successful electroporation and slice culture preparation12 of mouse brains, as well as time-lapse confocal imaging of cortical slice cultures.

<table><tbody><tr><td>isoflurane</td><td>Abbott Laboratories&amp;nbsp;</td><td>506949</td><td>Forene</td></tr><tr><td>6-well plate</td><td>Corning</td><td>351146</td><td></td></tr><tr><td>12-well plate</td><td>Corning</td><td>351143</td><td></td></tr><tr><td>non-absorbable surgical suture</td><td>Ethicon</td><td>K890H</td><td>3/8 circle, 13 mm, taper point</td></tr><tr><td>Micro Adson Forceps</td><td>Fine Science Tools</td><td>11018-12</td><td>serrated, length: 12 cm</td></tr><tr><td>fine scissors</td><td>Fine Science Tools</td><td>14063-09</td><td>angled to side, length: 9 cm</td></tr><tr><td>Mathieu Needle Holder</td><td>Fine Science Tools</td><td>12510-14</td><td>tungsten carbide, length: 14 cm</td></tr><tr><td>fine tipped forceps</td><td>Fine Science Tools</td><td>11370-40</td><td>straight, 11 cm</td></tr><tr><td>Vannas T&amp;uuml;bingen Spring Scissors</td><td>Fine Science Tools</td><td>15005-08</td><td>angled up, 9.5 cm</td></tr><tr><td>ring forceps</td><td>Fine Science Tools</td><td>11103-09</td><td>OD: 3mm, ID, 2.2 mm, length: 9 cm</td></tr><tr><td>HBSS (10X)</td><td>Gibco</td><td>14180046</td><td></td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030081</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Gibco</td><td>15140122</td><td></td></tr><tr><td>horse serum</td><td>Gibco</td><td>26050088</td><td></td></tr><tr><td>BME</td><td>Gibco</td><td>41010026</td><td></td></tr><tr><td>borosilicate glass capillaries</td><td>Harvard Apparatus</td><td>30-0016</td><td>1.0 OD x 0.58 ID x 100 L mm</td></tr><tr><td>anesthsesia system</td><td>Harvard Apparaus</td><td>72-6471</td><td></td></tr><tr><td>anesthetizing chamber</td><td>Harvard Apparaus</td><td>34-0460</td><td></td></tr><tr><td>fluosorber filter canister</td><td>Harvard Apparaus</td><td>34-0415</td><td></td></tr><tr><td>low melting point agarose</td><td>Invitrogen</td><td>16520100</td><td></td></tr><tr><td>vibrating blade microtome</td><td>Leica</td><td>VT1200 S</td><td></td></tr><tr><td>fluorescence stereo microscope</td><td>Leica</td><td>M205 FA</td><td></td></tr><tr><td>stereo microscope</td><td>Leica</td><td>M125</td><td></td></tr><tr><td>inverted fluorescence tissue culture microscope</td><td>Leica</td><td>DM IL LED</td><td></td></tr><tr><td>confocal laser scanning microscope</td><td>Leica</td><td>TCS SP5II</td><td></td></tr><tr><td>hybrid detector</td><td>Leica</td><td>HyD</td><td></td></tr><tr><td>objective, 40x/0.60 NA</td><td>Leica</td><td>11506201</td><td></td></tr><tr><td>microscope temperature control system</td><td>Life Imaging Services</td><td>Cube, Brick &amp;amp; Box</td><td></td></tr><tr><td>cell culture insert</td><td>Millipore</td><td>PICM0RG50</td><td></td></tr><tr><td>microgrinder</td><td>Narishige</td><td>EG-45</td><td>use 38&amp;deg; angle for beveling</td></tr><tr><td>microinjector</td><td>Parker Hannifin&amp;nbsp;</td><td>052-0500-900</td><td>Picospritzer III</td></tr><tr><td>carprofen</td><td>Pfizer Animal Health</td><td>NDC 61106-8507</td><td>Rimadyl</td></tr><tr><td>emdedding mold</td><td>Polysciences</td><td>18986-1</td><td></td></tr><tr><td>endotoxin-free plasmid maxi kit</td><td>Qiagen</td><td>12362</td><td></td></tr><tr><td>fast green</td><td>Sigma</td><td>F7252</td><td></td></tr><tr><td>laminin</td><td>Sigma</td><td>L2020</td><td></td></tr><tr><td>poly-L-lysine</td><td>Sigma</td><td>P5899</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td></td></tr><tr><td>D-glucose</td><td>Sigma</td><td>G6152</td><td></td></tr><tr><td>calcium chloride</td><td>Sigma</td><td>C7902</td><td></td></tr><tr><td>magensium sulfate</td><td>Sigma</td><td>M2643</td><td></td></tr><tr><td>sodium bicarbonate</td><td>Sigma</td><td>S6297</td><td></td></tr><tr><td>square wave electroporator</td><td>Sonidel</td><td>CUY21EDIT</td><td></td></tr><tr><td>tweezers with 5 mm platinum disk electrodes</td><td>Sonidel</td><td>CUY650P5</td><td></td></tr><tr><td>micropipette puller</td><td>Sutter Instrument</td><td>P-97</td><td></td></tr><tr><td>box filament</td><td>Sutter Instrument</td><td>FB255B</td><td>2.5 mm x 2.5 mm</td></tr><tr><td>micro-spoon spatula</td><td>VWR</td><td>231-0191</td><td>185 mm x 5 mm</td></tr><tr><td>glass bottom dish, 50 mm</td><td>World Precision Instruments</td><td>FD5040-100</td><td></td></tr></tbody></table>

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.

Previously, we have shown that genetic deletion of Bcl11a by in utero electroporation impairs radial migration of late-born upper-layer projection neurons10. Electroporation of a DNA plasmid vector containing Cre-IRES-GFP efficiently deleted Bcl11a in conditional Bcl11aflox/flox brains11. When we analyzed E14.5 electroporated brains three days after the electroporation, most control neurons had migrated into the CP, whereas many Bcl11a mutant neurons were stalled along the migratory route in the IZ and VZ/SVZ. Moreover, we found an increase in the number of multipolar cells at the expense of bipolar cells specifically in the IZ upon Bcl11a deletion suggesting defects in polarization10.
To directly and dynamically analyze migratory behavior of Bcl11a mutant neurons we used time-lapse confocal imaging of migrating neurons in organotypic slice culture prepared from electroporated brains. Overview time-lapse series using a 10X objective with a numerical aperture of 0.4 confirmed our results, that Bcl11a mutant projection neurons have defects in radial migration. Within 36 h many control neurons had migrated into the CP, whereas only few Bcl11a mutant neurons had left the IZ and migrated into the CP. Interestingly, it seemed that many Bcl11a mutant neurons did not migrate directly towards the pial surface of the brain, but slowed down migration and turned tangentially or even back towards the ventricle (Movie 1). To further analyze these findings we generated time-lapse series at a higher magnification using a 40x objective with a numerical aperture of 0.6. Our data show that Bcl11a mutant neurons failed to efficiently polarize and to continue radially oriented migration into the CP. Within 12 h, many Bcl11a mutant neurons failed to switch from a multipolar to a bipolar morphology and instead projected and retracted many processes into the surrounding environment (Movie 2, Figure 1A).
Furthermore, we traced the migration paths of individual neurons in different time-lapse series and used these to calculate specific parameters of migrating neurons. We found that Bcl11a mutant neurons repetitively undergo phases of several hour duration with reduced migration speed and random-like orientation changes (Movie 3; Figure 1B, red arrowheads). In particular, the speed profile of Bcl11a mutant neurons was shifted from higher speed (25 &#181;m/h) towards lower speed (5 &#181;m/h) in comparison to control neurons (Figure 1C). In line with this, the overall migration speed of Bcl11a mutant neurons was significantly reduced from 10.34 &#177; 0.34 &#181;m/h in controls to 6 &#177; 0.54 &#181;m/h (p = 2.4889E-05) (Figure 1D). Finally, the deviation from directed migration towards the pial surface of the brain was significantly increased from 17.15 &#177; 2.13&#176; in control to 40.16 &#177; 4.42&#176; in Bcl11a mutant neurons (p = 0.0002) (Figure 1E). Together these results demonstrate that time-lapse confocal imaging of GFP electroporated neurons in organotypic slice culture is a valuable method to study molecular mechanisms controlling the process of neuronal migration.
<img alt="Movie 1" src="/files/ftp_upload/55886/55886mov1.jpg" /> 
Movie 1: Migration of GFP Labeled Control in Comparison to Bcl11a Mutant Neurons in Cortical Slices. Bcl11aflox/flox brains were electroporated at E14.5 with a plasmid vector containing IRES-GFP (A) or Cre-IRES-GFP (B) and slice cultures were prepared at E16.5. VZ/SVZ; ventricular/subventricular zones; IZ, intermediate zone; CP, cortical plate. Scale bar = 100 &#181;m. The movie is comprised of 108 frames at the rate of 5 frames/s. Reprinted from Wiegreffe et al.10 with permission from Elsevier. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55886/Movie_1.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 2" src="/files/ftp_upload/55886/55886mov2.jpg" /> 
Movie 2: Polarization of a GFP Labeled Control in Comparison to Bcl11a Mutant Neurons in Cortical Slices. Bcl11aflox/flox brains were electroporated at E14.5 with a plasmid vector containing IRES-GFP (A) or Cre-IRES-GFP (B) and slice cultures were prepared at E16.5. Scale bar = 10 &#181;m. The movie is comprised of 25 frames at the rate of 5 frames/s. Reprinted from Wiegreffe et al.10 with permission from Elsevier. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55886/Movie_2.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 3" src="/files/ftp_upload/55886/55886mov3.jpg" /> 
Movie 3: Animated Traces with 1 h Interval Resolution of Representative Control and Bcl11a Mutant Neurons. Traces of migrating neurons were obtained from time-lapse series of E16.5 slice cultures of Bcl11aflox/flox brains that were electroporated at E14.5 with a plasmid vector containing IRES-GFP (A) or Cre-IRES-GFP (B). Scale bar = 20 &#181;m. Reprinted from Wiegreffe et al.10 with permission from Elsevier. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55886/Movie_3.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/55886/55886fig1.jpg" /> 
Figure 1: Migration Behavior of Bcl11a Mutant Upper-Layer Cortical Neurons. 
(A) Representative images of migrating GFP positive neurons in E16.5 slice cultures from Bcl11aflox/flox brains electroporated at E14.5 with either Cre-IRES-GFP or a control vector over a total imaging period of 12 h. (B) Representative traces with 1 hour interval resolution of migrating GFP positive neurons in E16.5 slice cultures from Bcl11aflox/flox brains electroporated at E14.5 with Cre-IRES-GFP or a control vector over total imaging periods of up to 22 h. Bcl11a mutant neurons frequently undergo repetitive phases of reduced migration speed and randomly changed orientation (marked by red arrowheads). (C) Speed profiles calculated from traces of migrating neurons as shown in B. (D-E) Quantification of speed (D) and deviation angle from radial orientation (E) of migrating GFP positive neurons in E16.5 slice cultures from Bcl11aflox/flox brains electroporated at E14.5 with Cre-IRES-GFP or a control vector (n = 15). Mean &#177; s.e.m.; Student&#39;s t test; ***p &#60;0.001. Scale bar = 10 &#956;m. Reprinted from Wiegreffe et al.10 with permission from Elsevier. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/55886/55886fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Time-lapse Confocal Imaging of Migrating Neurons in Organotypic Slice Culture of Embryonic Mouse Brain Using In Utero Electroporation at JoVE.com

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.

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 ...

time-lapse-confocal-imaging-migrating-neurons-organotypic-slice

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

10.2K Views.  Ulm University. 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.

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

Organotypic Slice Culture of GFP-expressing Mouse Embryos for Real-time Imaging of Peripheral Nerve Outgrowth

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in which the enhanced version of the green fluorescent protein (EGFP) is exclusively expressed in all neurons of the developing central and peripheral nervous system1, allowing the possibility to both film the innervation of the forelimb and to manipulate this process with pharmacological and genetic techniques2. The most critical parameter in the successful cultivation of such slice cultures is the method by which the slices are prepared. After extensive testing of a variety of methods, we have found that a vibratome is the best possible device to slice the embryos such that they routinely result in a culture that demonstrates viability over a period of several days, and most importantly, develops in an age-specific manner. For mid-gestation embryos, this includes the normal outgrowth of spinal nerves from the spinal cord and the dorsal root ganglia to their targets in the periphery and the proper determination of skeletal and muscle tissue. 
In this work, we present a method for processing whole embryos of embryonic day (E) E10 to E12 into 300 - 400 micrometer slices for cultivation in a standard tissue culture incubator, which can be studied for up to two days after slice preparation. Critical for the success of this approach is the use of a vibratome to slice each agarose-embedded embryo. This is followed by the cultivation of the slices upon Millicell culture membrane inserts placed upon a small volume of medium, resulting in an interface culture technique. One litter with an average of 7 embryos routinely produces at least 14 slices (2-3 slices of the forelimb region per embryo), which varies slightly due to the age of the embryos as well as to the thickness of the slices. About 80% of the cultured slices show nerve outgrowth, which can be measured througout the culturing period2. Representative results using the tauGFP mouse line are demonstrated.

We present a method to prepare organotypic slices of mid-gestation mouse embryos for the cultivation and time-lapse imaging of peripheral nerve outgrowth.

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in ...

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 ...

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the first born neurons remain closer to the ventricle while the last born neurons migrate past the first born neurons towards the surface of the brain1. In addition to neuronal migration2, a key process for normal cortical function is the regulation of neuronal morphogenesis3. While neuronal morphogenesis can be studied in vitro in primary cultures, there is much to be learned from how these processes are regulated in tissue environments.
We describe techniques to analyze neuronal migration and/or morphogenesis in organotypic slices of the cerebral cortex4,6. A pSilencer modified vector is used which contains both a U6 promoter that drives the double stranded hairpin RNA and a separate expression cassette that encodes GFP protein driven by a CMV promoter7-9. Our approach allows for the rapid assessment of defects in neurite outgrowth upon specific knockdown of candidate genes and has been successfully used in a screen for regulators of neurite outgrowth8. Because only a subset of cells will express the RNAi constructs, the organotypic slices allow for a mosaic analysis of the potential phenotypes. Moreover, because this analysis is done in a near approximation of the in vivo environment, it provides a low cost and rapid alternative to the generation of transgenic or knockout animals for genes of unknown cortical function. Finally, in comparison with in vivo electroporation technology, the success of ex vivo electroporation experiments is not dependant upon proficient surgery skill development and can be performed with a shorter training time and skill.

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

To conduct a rapid assessment of the function of genes in the development of cerebral cortex, we describe methods involving the ex vivo electroporation of plasmids co-expressing inhibitory RNA (RNAi) and GFP in murine embryonic cortex. This protocol is amenable to the study of various aspects of neurodevelopment such as neurogenesis, neuronal migration and neuronal morphogenesis including dendrite and axon outgrowth.

The cerebral cortex directs higher cognitive functions. This six layered structure is generated in an inside-first, outside-last manner, in which the ...

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 ...

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 ...

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate tangentially from their place of origin in the ventral telencephalon (including from the medial and caudal ganglionic eminences (MGE, CGE)) to the dorsal cortical plate in response to a variety of intrinsic and extrinsic cues. Different methodologies have been developed over the years to genetically manipulate specific pathways and investigate how they regulate the dynamic cytoskeletal changes required for proper IN migration. In utero electroporation has been extensively used to study the effect of gene repression or overexpression in specific IN subtypes while assessing the impact on morphology and final position. However, while this approach is readily used to modify radially migrating pyramidal cells, it is more technically challenging when targeting INs. In utero electroporation generates a low yield given the decreased survival rates of pups when electroporation is conducted before e14.5, as is customary when studying MGE-derived INs. In an alternative approach, MGE explants provide easy access to the MGE and facilitate the imaging of genetically modified INs. However, in these explants, INs migrate into an artificial matrix, devoid of endogenous guidance cues and thalamic inputs. This prompted us to optimize a method where INs can migrate in a more naturalistic environment, while circumventing the technical challenges of in utero approaches. In this paper, we describe the combination of ex utero electroporation of embryonic mouse brains followed by organotypic slice cultures to readily track, image and reconstruct genetically modified INs migrating along their natural paths in response to endogenous cues. This approach allows for both the quantification of the dynamic aspects of IN migration with time-lapse confocal imaging, as well as the detailed analysis of various morphological parameters using neuronal reconstructions on fixed immunolabeled tissue.

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

Here, we provide a low-cost and reliable method to generate electroporated brain organotypic slice cultures from mouse embryos suitable for confocal microscopy and live-imaging techniques.

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate ...

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones -the sensory structures located at the distal tips of developing axons- execute this process. Studying the structure and dynamics of the growth cone is crucial to understanding axonal development and the interactions with the surrounding central nervous system (CNS) that enable it to form neural circuits. This is essential when devising methods to reintegrate axons into neural circuits following injury in fundamental research and pre-clinical contexts. Thus far, the general understanding of growth cone dynamics is primarily founded on studies of neurons cultured in two dimensions (2D). Although undoubtedly fundamental to the current knowledge of growth cone structural dynamics and response to stimuli, 2D studies misrepresent the physiological three-dimensional (3D) environment encountered by neuronal growth cones in intact CNS tissue. More recently, collagen gels were employed to overcome some of these limitations, enabling the investigation of neuronal development in 3D. However, both synthetic 2D and 3D environments lack signaling cues within CNS tissue, which direct the extension and pathfinding of developing axons. This protocol provides a method for studying axons and growth cones using organotypic brain slices, where developing axons encounter physiologically relevant physical and chemical cues. By combining fine-tuned in utero and ex utero electroporation to sparsely deliver fluorescent reporters along with super-resolution microscopy, this protocol presents a methodological pipeline for the visualization of axon and growth cone dynamics in situ. Furthermore, a detailed toolkit description of the analysis of long-term and live-cell imaging data is included.

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

This protocol demonstrates a straightforward and robust method to study in situ axon growth and growth cone dynamics. It describes how to prepare ex vivo physiologically relevant acute brain slices and provides a user-friendly analysis pipeline.

During neuronal development, axons navigate the cortical environment to reach their final destinations and establish synaptic connections. Growth cones ...

Visualizing Neuronal and Glial Cell Dynamics in the Developing Mouse Brain

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.

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.

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

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Chapters in this video

Organotypic Slice Culture of GFP-expressing Mouse Embryos for Real-time Imaging of Peripheral Nerve Outgrowth

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

Methods for Study of Neuronal Morphogenesis: Ex vivo RNAi Electroporation in Embryonic Murine Cerebral Cortex

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

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

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

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