This work was supported by operating grants from the Savoy Foundation and the CURE Epilepsy Foundation and by equipment grants from the Canadian Foundation for Innovation to E.R (confocal microscope) and G.H (spinning disk confocal microscope). E.R. receives a career award from the Fonds de recherche du Qu&#233;bec-Sant&#233; (FRQ-S; Clinician-scientist Award) as well as from the Canadian Institutes for Health Research (CIHR; Young Investigator Award). G.H. is a senior scholar of the FRQ-S. L.E is the recipient of the Steriade-Savoy postdoctoral training award from the Savoy Foundation, the CHU Sainte-Justine Foundation postdoctoral training award and the FRQ-S postdoctoral training award, in partnership with the Foundation of Stars. This project has been made possible by Brain Canada through the Canada Brain Research Fund, with the financial support of Health Canada, awarded to L.E.

All experiments involving animals were approved by the Comit&#233; Institutionnel des Bonnes Pratiques avec les Animaux de Recherche (CIBPAR) at the CHU Sainte-Justine Research Center and were conducted in accordance with the Canadian Council on Animal Care&#39;s guide to the Care and Use of Experimental Animals (Ed. 2).
The protocol described here was optimized for electroporation of embryos at embryonic day (e) 13.5, at a time when MGE-derived INs are actively generated, before the peak of CGE-derived INs production45,46. Furthermore, to bias the electroporation towards GABAergic INs, we use a promoter selectively expressed in INs (for instance the Dlx5/6 promoter with its minimal enhancer)47.
1. Preparation of Solutions for Electroporation and Organotypic Slice Cultures
<ol>
	<li>Prepare 125 mL of sterile culture medium.
	<ol>
		<li>Measure 125 mL of regular neuron-specific culture medium (see Table of Materials for formulation) in a sterilized bottle in a previously UV-sterilized biosafety cabinet sprayed with 70% ethanol. Add 2.25 mL of 50x serum-free neuron-specific supplement, 1.75 mL of 200mM glutamine (final concentration of 0.5 mM) and 6.25 mL of heat-inactivated horse serum previously aliquoted under sterile conditions. Mix thoroughly, aliquot in 15 mL sterile conical tubes, and store at 4 &#176;C. 
		NOTE: Prepared culture medium can be stored for up to 3 weeks at 4 &#176;C.</li>
		<li>Divide a 100X stock solution of Botteinstein&#39;s N-2 formulation48 into 150 &#181;L aliquots under sterile conditions and freeze at -20&#176;C until use.</li>
	</ol>
	</li>
	<li>Prepare 1 L of sterile artificial cerebrospinal fluid (ACSF).
	<ol>
		<li>Measure 800 mL of distilled water in a 1 L beaker. Add 25.67 g of sucrose, 5.08 g of sodium chloride (NaCl), 2.18 g of sodium bicarbonate (NaHCO3), 1.80 g of glucose, 0.19 g of potassium chloride (KCl), 0.15 g of monobasic anhydrous sodium phosphate (NaH2PO4), 1 mL of 1M stock CaCl2.2H2O and 2 mL of 1M stock MgSO4.7H2O. Stir to dissolve at room temperature. Add distilled water to reach a total volume of 1 L.</li>
		<li>Using a 0.22 &#181;m filter, filter the solution into a sterile bottle in a sterilized biosafety cabinet and store at 4 &#176;C for up to 1 month.</li>
	</ol>
	</li>
	<li>Prepare a fresh solution of 4% agarose in ACSF before each experiment.
	<ol>
		<li>Measure 25 mL of previously prepared ACSF in a 50-mL sterile conical tube and add 1 g of low-melting point agarose.</li>
		<li>Heat for 45 s in a microwave oven. To avoid spilling, interrupt heating every 3 - 4 s when boiling starts, open the tube to let the pressure out, close it again and agitate manually to mix the agarose. Repeat until the agarose solution is homogeneous. Keep the agarose solution at 42 &#176;C during the remainder of the experiment to avoid solidification. 
		NOTE: Higher temperatures will damage brain tissues.</li>
	</ol>
	</li>
</ol>
2. Preparation of Plasmids for Injection
<ol>
	<li>Pull a glass microinjection pipette
	<ol>
		<li>Set the micropipette puller with the proper parameters, secure the glass capillary in the provided space and make sure it is centered with the filament.</li>
		<li>Press the pull button.</li>
		<li>Carefully remove the newly made microinjection pipettes from the heat puller and place in a box or clean petri dish until further use to avoid damaging the tip.</li>
	</ol>
	</li>
	<li>Set-up the biosafety cabinet with all the instruments needed for this experiment (see Figure 1A), generously spray all instruments in the biosafety cabinet with 70% ethanol and sterilize the instruments and environment with UV light for 15 - 20 min.</li>
	<li>During the sterilization step, thaw plasmids on ice (4 &#176;C).</li>
	<li>Measure 10 &#181;L of the plasmid from a stock solution (4 &#181;g/&#181;L) into a clean 1.5 mL centrifuge tube. Add 0.01% of Fast Green FCF. Mix gently, spin briefly and keep on ice until use. 
	NOTE: Maxi-prep DNA should be prepared according to the manufacturer&#39;s protocol using an endotoxin-free maxi-prep kit. DNA can be solubilized in TE buffer or nuclease-free H2O and the preparation of the plasmid with dye solution can, but does not need to be performed under sterile conditions. Plasmids from Maxi-prep should be aliquoted to avoid multiple freeze-thaw cycles. Aliquoted plasmids should be mixed with the dye no more that 2 h before use and should not be refrozen for further use.</li>
	<li>After sterilization, prepare the nano-injector as follows.
	<ol>
		<li>Choose one of the previously prepared glass microinjection pipette stored in a clean box or petri dish and use small tweezers to cut the tip of the pipette in a beveled way to achieve an outer diameter of roughly 15 &#181;m. 
		NOTE: The outer diameter given here is an approximate measure to give the user an idea of what we use in our experiments and has been optimized in order to facilitate the piercing of the skull for plasmid injection without damaging the brain, while allowing for the fluid loading and release of the DNA. The outer diameter can be measured by looking at the cut tip of the glass microinjection pipette apposed to a micrometric scale bar under a bright field microscope.</li>
		<li>Use a syringe to fill the micropipette with mineral oil from its unpulled end (to expel all the air).</li>
		<li>Insert the filled glass micropipette in the nano-injector by following the manufacturer&#39;s instructions.</li>
		<li>Empty 2/3rds of the glass micropipette (keeping enough oil to prevent air entry).</li>
	</ol>
	</li>
	<li>Carefully insert the prepared micropipette in the tube containing the plasmid/dye solution and fill the glass micropipette with the plasmid/dye solution.</li>
</ol>
3. Collection of Mouse Embryos from Pregnant Females
<ol>
	<li>Monitor breeding females daily to assess for vaginal plugging, preferably at the same time daily (early afternoon). Day e0.5 corresponds to the first day when a vaginal plug is observed. 
	NOTE: The experiments described here can be conducted in wild-type mice. However, to facilitate the identification of the MGE and to label all GABAergic INs (or specific sub-sets such as MGE-derived INs), transgenic animals can be used (e.g.: GAD67EGFP; Dlx5/6Cre with a Cre-reporter allele, etc.47,49). In this situation, the experimental plasmid injected should express another fluorophore (for instance mCherry or TdTomato) to allow visualization of the transfected INs (yellow) that can be compared with non-transfected INs (green).</li>
	<li>Sacrifice the female at embryonic day e13.5, by neck dislocation. 
	NOTE: Anesthetic agents given at the time of sacrifice may impact IN migration and survival50,51 and should be avoided.</li>
	<li>Collect embryos by C-section as follows.
	<ol>
		<li>Generously spray the female abdomen with 70% ethanol. Pull the abdominal skin up with a pair of sterilized forceps and, with the other hand, use sterilized surgical scissors to cut the skin from the abdomen.</li>
		<li>With a second pair of sterilized forceps and scissors, pull the abdominal fascia up and cut it while carefully avoiding the uterus.</li>
		<li>Using a third pair of sterilized forceps and scissors, pull the uterine horns and cut them out of the pelvic cavity. Place the dissected uterine horns in a sterile 60-mm petri dish filled with neural-based culture medium supplemented with amino acids, vitamins and inorganic salts (see Table of Materials for commercially available product).</li>
	</ol>
	</li>
	<li>In a sterile biosafety cabinet, use two pairs of fine tweezers (one in each hand) to dissect the embryos out of the placenta and isolate the heads by decapitation.</li>
	<li>Bevel-cut the tip of a sterile 3 mL plastic transfer pipette, aspirate the heads and transfer them in a new sterile 60-mm petri dish layered with solidified black wax and filled with the same neural-based supplemented culture medium as above. 
	NOTE: This step minimizes the transfer of contaminants (mouse hair, blood). The black wax is used to stabilize the head during dissection. Culture media do not need to be oxygenated during these procedures.</li>
</ol>
4. Intraventricular Plasmid Injections and Ex Vivo Electroporation of the MGE
NOTE: The following steps must be performed under sterile conditions in the previously prepared biosafety cabinet.
<ol>
	<li>Place the 60-mm petri dish layered with black wax and containing the decapitated heads in neural-based supplemented culture medium under the binoculars in the biosafety cabinet.</li>
	<li>Stabilize the head, the rostral part facing right, with fine tweezers with the left hand and use the nano-injector in the right hand to inject 1 - 2 &#181;L of the plasmid/dye solution into the right lateral ventricle. 
	NOTE: Co-expression experiments can be conducted by co-electroporating a rescue plasmid and a shRNA-expressing plasmid by mixing both plasmids at equimolar concentrations.</li>
	<li>Electroporate the injected brain.
	<ol>
		<li>Place the head between the electrodes with the negative electrode positioned dorsally and parallel to the head and the positive electrode towards the ventral side of the head to target the MGE.</li>
		<li>Once the electrodes are well positioned, deliver 4 square pulses of 40 V for 50 ms at 500 ms inter-pulse intervals.</li>
		<li>Remove any residual tissue from the electrodes using tweezers already placed in the biosafety cabinet. 
		NOTE: These parameters have been optimized specifically for the electroporator used in our experiments. We recommend that the users perform optimizing tests beforehand if using a different type of electroporator.</li>
		<li>Repeat steps 4.1 to 4.3 for all remaining brains. 
		NOTE: Although this protocol describes the manipulations required for one brain, it is possible to inject up to 4 brains sequentially before electroporating each brain, thus increasing the yield. This strategy is especially advantageous when 2 or more different plasmids are injected sequentially (e.g. control or experimental plasmid) during the same experiment (allowing for comparison between littermates). In addition, it is possible to inject and electroporate simultaneously both sides of the brain to increase the yield, by positioning the electrodes completely parallel to the brain surface.</li>
	</ol>
	</li>
</ol>
5. Brain Dissection and Organotypic Slice Cultures
<ol>
	<li>While still manipulating in the sterile environment of the biosafety cabinet, dissect the brain out of the skull.
	<ol>
		<li>Stabilize the head on the layer of black wax by inserting a needle into each eye while carefully avoiding the brain.</li>
		<li>Use a pair of fine tweezers to hold the left side of the neck and a second pair of fine tweezers to tear the skin from the skull, from back to front.</li>
		<li>While holding the head laterally with tweezers in one hand, use another pair of tweezers in the other hand to carefully cut the skull at the level of the brainstem and gently pull the skull up. With each tweezer, cut the skull in the sagittal plane (midline) towards the front, and then incise laterally to liberate the skull fragments.</li>
		<li>Lift the brainstem and carefully cut the meninges and cranial nerves until the brain is completely out of the skull. 
		NOTE: All steps described in 5.1 should be performed under stringent sterile conditions in a biosafety cabinet.</li>
	</ol>
	</li>
	<li>Embed the brain in 4% low-melting point agarose for sectioning.
	<ol>
		<li>Fill a 35-mm petri dish with the agarose solution prepared above (kept liquid at 42 &#176;C).</li>
		<li>Quickly transfer an electroporated brain to the agarose-filled dish using the previously cut transfer pipette. Keep the dish at room temperature.</li>
		<li>Stir the agarose with a metal stick to keep the brain in the middle of the well (to prevent sinking) and position the brain in a rostro-caudal plane parallel to the dish. Stop stirring when the agarose starts to solidify, to avoid any brain damage.</li>
		<li>Use a razor blade to cut the agarose surrounding the brain in order to form a rectangular block, leaving a margin of 1 - 2 millimeters around the brain. Ensure that the rostral part of the brain is perpendicular to the anterior limit of the block to facilitate orientation for the subsequent sectioning steps.</li>
		<li>Repeat for each brain. 
		NOTE: It is possible to cut more than one brain at a time (maximum 3) by shaping separate agarose blocks while setting each brain at different heights.</li>
	</ol>
	</li>
	<li>Vibratome coronal sections and slice culture.
	<ol>
		<li>Thaw one aliquot of 100X N-2 supplement (150 &#181;L) on ice and add to 15 mL aliquoted culture medium under sterile conditions.</li>
		<li>Transfer 750 &#181;L of culture medium (with 1X N2 supplement) to each well of a 6-well culture plate.</li>
		<li>With curved tweezers, place one cell culture insert (30 mm diameter, 0.4 &#181;m pore size, PTFE) in each medium-filled well.</li>
		<li>Fill the vibratome bath with continuously oxygenated ACSF. Cool to 4 &#176;C with ice surrounding the bath, or use a refrigerated vibratome.</li>
		<li>Set the vibratome speed to 0.150 mm/s and the frequency to 80 Hertz.</li>
		<li>Glue the agarose block on the vibratome platform, rostral edge facing down and ventral edge facing the user.</li>
		<li>Cut the brain in coronal sections to obtain 250 &#181;m-thick sections (at 4 &#176;C).</li>
		<li>With sterilized spatulas, collect 2 - 3 sections containing the MGE and place all brain sections from one animal on a single 30-mm membrane insert, while carefully avoiding overlap between sections. Place the insert in one well of a 6-well culture plate (containing 750 &#181;L of supplemented culture medium, as described above). Alternatively, each section can be placed on a separate 13-mm diameter membrane in a 12-well culture plate filled with 500 &#181;l supplemented culture medium. The amount of culture medium recommended for each well allows the brain sections to be nourished by the media without being submerged. 
		NOTE: The steps described in 5.3.6 and 5.3.7 are not carried out under complete sterile conditions, unless the vibratome can be sterilized and used in a biosafety cabinet. Therefore, it is crucial to conduct these steps carefully to avoid any contamination. Appropriate protective equipment (clean mask, surgical gloves and lab coat) should be worn at all times and body parts, even covered, such as hair, face and hands, should never pass over culture plates (with or without culture medium). It is also recommended to spray 70% ethanol frequently on the gloves and spatulas used to collect the brain sections.</li>
		<li>Place the culture plate in a ventilated sterile incubator at 37 &#176;C with 60% humidity and 5% CO2 for 48 or 72 h. 
		NOTE: These incubation times were optimized for time-lapse imaging of MGE-derived INs and confocal imaging of fixed slices, respectively. Optimal incubation times should be tested beforehand for each experimental design. In addition, if the chosen incubation is 72 h and under, there is no need to change the culture medium. For longer incubation times, the culture medium should be changed every 2 - 3 days.</li>
		<li>After the desired incubation time, transfer the sections of interest to an 8-chambered coverslip and add 3 - 5 &#181;L of culture medium. Place the coverslip in an environmental chamber (37 &#176;C, 60% humidity, 5% CO2) connected to an inverted spinning disk confocal equipped with a computer-assisted acquisition software to set-up the time-lapse imaging session. 
		NOTE: Alternatively, sections can be fixed with 4% paraformaldehyde (overnight at 4 &#176;C or 2 h at room temperature) and subsequently immunostained with different antibodies for visualization of the morphological features of electroporated INs under a confocal microscope. Although eGFP and mCherry can be visualized by confocal microscopy without any counter-staining procedure, we recommend performing immunohistochemistry against GFP and mCherry to enhance the signal since the fixation process can reduce fluorescence, reducing the detection of finer components of embryonic neurons, such as smaller branches in the leading or trailing processes.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose. The views expressed herein do not necessarily represent the views of the Minister of Health or the Government of Canada.

In this article, we provide a reliable method for performing ex utero electroporation of the mouse MGE at e13.5 and for the generation of organotypic cultures of embryonic brain slices. Although in vitro methods, such as the Boyden Chamber Assay, are relatively easy to perform and can be used to assess the specific roles of different genes and proteins without the interference of other factors, they preclude the investigation of IN migration dynamics with regards to directionality and migration path<sup...

Cortical GABAergic interneurons (INs) are diverse with regards to their biochemical properties, physiological properties and connectivity, and they mediate different functions in mature networks1,2,3,4,5. The specification of different subtypes of cortical INs is tightly regulated through genetic cascades that have been extensively studied1,2. The majority (70%) of cortical GABAergic INs originate from progenitors in the medial ganglionic eminence (MGE), a ventrally located embryonic structure, and must migrate across relatively long distances to reach the cortical plate1,2,6. While cortical pyramidal cells migrate radially from the ventricular zone (VZ) to the cortical plate along the radial glia scaffold, the tangential migration of INs, which are not attached to such a scaffold, requires a variety of intrinsic and extrinsic cues to attract migrating neurons towards the cortical plate, while guiding them away from non-cortical structures2,7,8. After exiting the cell cycle, INs are repelled from the MGE by chemo-repulsive cues expressed within the VZ of the MGE, which triggers tangential migration towards the cortical plate9,10. Migrating INs avoid the striatum with the aid of different repulsive cues11 and, after reaching the cortical plate, they switch from a tangential to a radial migration mode and reach their final laminar position, partly in response to cues from pyramidal cells12 and other cellular populations13. The migration of INs, as for other neuronal populations, involves various dynamic morphological changes to permit the actual movement of the neuron. This so-called neuronal locomotion is characterized by repetitive cycles of three successive steps: the elongation of a leading process, an active anterograde motion of the nucleus (nucleokinesis), and the retraction of the trailing process14. IN migration is regulated by numerous intrinsic and extrinsic cues that drive the branching and active remodeling of the leading process to guide INs in the proper direction, determining both orientation and speed of migration14,15,16.
The determinants regulating cortical IN migration have been extensively studied in recent years1,2,7,17,18,19,20, and disruption in some of these molecular actors has been postulated to lead to neurodevelopmental disorders, such as pediatric refractory epilepsy or autism spectrum disorders1,2,21,22,23,24. Therefore, the development of various in vitro and in vivo approaches has been pursued to significantly advance our ability to study this dynamic process, as previously reviewed25. In vitro methods, including the Boyden chamber assay and the Stripe Choice Assay, provide the fastest and most reproducible means of assessing the requirement and cell-autonomous impact of specific genes or proteins during neuronal migration, without the influence of other factors25. These assays are particularly useful when combined with live-imaging8,26,27. With these techniques, INs are easily retrieved from e13.5 MGE and isolated by enzymatic and mechanical dissociation, after which different signaling pathways and guidance cues can be investigated, as illustrated previously8,28. However, these assays take place in an artificial extracellular matrix in the absence of three-dimensional tissue architecture, which may alter neuronal behavior and cell properties, potentially affecting cell migration and/or survival25. To circumvent these limitations, ex vivo MGE explants have been developed as an alternative tool to quantify the dynamic morphological changes occurring during migration along with parameters such as speed and orientation14,29. Generating MGE explants is relatively straightforward and has been extensively described elsewhere30. It entails the plating of a small extract of the MGE on a monolayer of mixed cortical cells or in a mixture of matrigel and collagen in the presence of attractive or repulsive cues25, although the latter are optional31. MGE explants allow for high resolution imaging of sparsely labeled cells, simplifying the study of intracellular processes, such as cytoskeletal remodeling during leading process branching, as shown previously32,33,34 and in the present study. MGE explants have been used successfully to assess dynamic cytoskeletal changes during IN migration in a 2D environment, for instance after specific pharmacological or chemotactic manipulations (see, for example, Tielens et al. 201633). However, with this approach, INs migrate within an artificial matrix, and this might alter IN behavior and the reproducibility and significance of the experimental results.
By contrast, in utero electroporation enables the genetic manipulation of INs in their native environment and is a widely used method to rapidly and efficiently assess the impact of gain and loss of gene function while circumventing the limitations of costly and time-consuming knockout and knock-in strategies25,35. In utero electroporation can be biased towards IN progenitors by using cell type specific promoters and by positioning the electrodes towards ventromedial structures, including the MGE36. Furthermore, in utero electroporation allows for the timely expression of experimental constructs within 1 - 2 days, as compared to the 7 - 10 days required for construct expression using viral vectors25. However, in utero electroporation of IN progenitors tends to be low-yield. Indeed, although pyramidal cell progenitors located in the dorsal ventricular zone can be efficiently transfected using in utero electroporation, targeting more ventrally located structures, such as the MGE, is more technically challenging, especially in small e13.5 embryos, and the high rate of embryonic lethality further reduces the experimental yield25.
To circumvent some of the technical limitations associated with in vitro MGE explant experiments and in vivo in utero electroporation, ex vivo organotypic slice cultures have been developed8,37,38,39. Brain organotypic slice cultures offer the advantage of mimicking in vivo conditions, while being less expensive and time-consuming than generating animal models25. Indeed, these preparations allow an easy access to the MGE, along with the specific visualization of INs, and can be combined with focal electroporation to investigate specific molecular pathways in INs migrating in a more physiological environment8,39,40,41. We have therefore optimized an approach for organotypic cultures38, which we combined with ex utero electroporation and time-lapse confocal imaging, to further assess the morphological and dynamic process occurring during tangential migration of MGE-INs. The present protocol was adapted and optimized from others who have used ex utero or in utero brain electroporation and organotypic slice cultures to study the migration of pyramidal cells42,43 and cortical INs36,39,44. Specifically, mouse embryos are decapitated and the MGE is electroporated ex vivo after the intraventricular injection of the experimental plasmids, allowing more efficient and precise targeting of MGE progenitors than what can be achieved with in utero electroporation. The brains are then extracted and sectioned into whole brain coronal slices that can be cultured for a few days, thus allowing continuous tracking and imaging of transfected INs. This approach typically labels 5 - 20 tangentially migrating INs per brain slice, minimizing the number of experimental iterations required to reach statistical significance, while labeling a sufficiently sparse neuronal population to ensure easy separation of individual neurons for reconstruction and fine morphological assessment. Furthermore, compared to MGE explants, organotypic cultures ensure that migrating INs are exposed to a more natural environment, including locally secreted chemokines and inputs from thalamic afferents. This approach is thus well suited to quantify the directionality and migratory path adopted by transfected INs, while offering sufficient anatomical details to allow the characterization of finer dynamic processes such as leading process branching, nucleokinesis and trailing process retraction.

<table>
	<tbody>
		<tr>
			<td>Neurobasal Medium</td>
			<td>ThermoFisher Scientific</td>
			<td>21103049</td>
			<td>Commercially available neuron-specific culture medium. Complete formulation available on this website: https://www.thermofisher.com/ca/en/home/technical-resources/media-formulation.251.html</td>
		</tr>
		<tr>
			<td>B-27 serum-free supplement</td>
			<td>ThermoFisher Scientific</td>
			<td>17504044</td>
			<td>50X Serum-free neuron specific supplement</td>
		</tr>
		<tr>
			<td>15 mL sterile centrifuge tubes</td>
			<td>Sarstedt</td>
			<td>62.554.002</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s (1X) L-15 Medium (+ L-Glutamine)</td>
			<td>ThermoFisher Scientific</td>
			<td>11415064</td>
			<td>Commercially available neural-based culture medium supplemented with amino acids, vitamins and inorganic salts. Complete formulation available on the distributor&#39;s website&#160;</td>
		</tr>
		<tr>
			<td>L-Glutamine</td>
			<td>Invitrogen</td>
			<td>25030-081</td>
			<td></td>
		</tr>
		<tr>
			<td>Horse serum, heat inactivated</td>
			<td>Millipore-Sigma</td>
			<td>H1138-500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Neurocell supplement N-2 100X</td>
			<td>Wisent</td>
			<td>305-016</td>
			<td>Botteinstein&#39;s N-2 Formulation</td>
		</tr>
		<tr>
			<td>VWR Square PETG Media Bottles 125 mL</td>
			<td>VWR</td>
			<td>89132-062</td>
			<td></td>
		</tr>
		<tr>
			<td>Class II Type A Biosafety Cabinet</td>
			<td>Nuaire</td>
			<td>NU-540</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>BioShop</td>
			<td>SUC700.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>BioShop</td>
			<td>SOD001.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>ThermoFisher Scientific</td>
			<td>S233-500</td>
			<td></td>
		</tr>
		<tr>
			<td>D+ glucose</td>
			<td>Millipore-Sigma</td>
			<td>G7528-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>ThermoFisher Scientific</td>
			<td>P217-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic anhydrous</td>
			<td>BioShop</td>
			<td>SPM400.500</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate</td>
			<td>BiosShop</td>
			<td>MAG522</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>BioShop</td>
			<td>AGA002.500</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL sterile centrifuge tubes</td>
			<td>Sarstedt</td>
			<td>62.547.004</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL centrifuge tubes</td>
			<td>Sarstedt</td>
			<td>72.690.001</td>
			<td></td>
		</tr>
		<tr>
			<td>P-97 Flaming/Brown Micropipette puller</td>
			<td>Sutter Instruments Co.</td>
			<td>Model P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>0.4 mm I.D. x 75 mm Capillary Tube</td>
			<td>Drummond scientific</td>
			<td>1-000-800/12</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>VWR</td>
			<td>E193</td>
			<td></td>
		</tr>
		<tr>
			<td>5 mL syringe</td>
			<td>Becton Dickinson &#38; Co</td>
			<td>309646</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral Oil (heavy)</td>
			<td>Rougier Pharma</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>WPI Swiss Tweezers #5</td>
			<td>World Precision Instruments</td>
			<td>504511</td>
			<td>11 cm, straight, 0.06x0.07mm tips, antimagnetic. You will need 2 of these.</td>
		</tr>
		<tr>
			<td>WPI Swiss Tweezers #7</td>
			<td>World Precision Instruments</td>
			<td>504504</td>
			<td>11.5 cm, 0.18x0.2mm, curved tips</td>
		</tr>
		<tr>
			<td>HTC Tweezers</td>
			<td>World Precision Instruments</td>
			<td>504617</td>
			<td>11 cm, Straight, flat</td>
		</tr>
		<tr>
			<td>Operating scissors</td>
			<td>World Precision Instruments</td>
			<td>501225</td>
			<td>16 cm, Sharp/sharp, straight. You will need 3 of these.</td>
		</tr>
		<tr>
			<td>Dressing Forceps</td>
			<td>World Precision Instruments</td>
			<td>501217</td>
			<td>12.5 cm, straight, serrated</td>
		</tr>
		<tr>
			<td>Iris Forceps</td>
			<td>World Precision Instruments</td>
			<td>504478</td>
			<td>10.2 cm, full curve, serrated</td>
		</tr>
		<tr>
			<td>DeBakey Tissue Forceps</td>
			<td>World Precision Instruments</td>
			<td>501996</td>
			<td>15 cm, 45&#176; angle, Delicate Jaw, 1.5mm wide</td>
		</tr>
		<tr>
			<td>Fisherbran Microspatula with rounded ends</td>
			<td>FisherScientific</td>
			<td>21-401-5</td>
			<td>You will need 2 of these.</td>
		</tr>
		<tr>
			<td>Nanoject II Auto-Nanoliter Injector</td>
			<td>Drummond scientific</td>
			<td>3-000-204</td>
			<td></td>
		</tr>
		<tr>
			<td>TC Dish 60, Standard</td>
			<td>Sarstedt</td>
			<td>83.3901</td>
			<td>60-mm dish</td>
		</tr>
		<tr>
			<td>Tissue culture dish</td>
			<td>Sarstedt</td>
			<td>83.1800</td>
			<td>35-mm dish</td>
		</tr>
		<tr>
			<td>Black Wax</td>
			<td>FisherScientific</td>
			<td>S17432</td>
			<td></td>
		</tr>
		<tr>
			<td>Transfer pipettes&#160;</td>
			<td>Ultident</td>
			<td>170-CTB700-212</td>
			<td>3 mL, small bulb</td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Leica Biosystems</td>
			<td>Leica M80</td>
			<td>In replacement to our stereomicroscope which has been discontinued by the manufacturer (StereoMaster from FisherScientific)</td>
		</tr>
		<tr>
			<td>Electro Square Porator</td>
			<td>BTX Harvard Apparatus</td>
			<td>ECM 830</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezertrodes, Plattinum Plated, 3mm</td>
			<td>BTX Harvard Apparatus</td>
			<td>45-0487</td>
			<td></td>
		</tr>
		<tr>
			<td>25G 1 1/2</td>
			<td>Becton Dickinson &#38; Co</td>
			<td>305127</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica VT1000S Vibrating blade microtome</td>
			<td>Leica Biosystems</td>
			<td>VT1000S</td>
			<td></td>
		</tr>
		<tr>
			<td>GEM, Single edge razor blade</td>
			<td>Electron Microscopy Sciences</td>
			<td>71952-10</td>
			<td>Remove the blunt end before inserting in the blade designated space of the vibratome</td>
		</tr>
		<tr>
			<td>&#181;-Slide 8 well</td>
			<td>Ibidi</td>
			<td>80827</td>
			<td>Pack of 15</td>
		</tr>
		<tr>
			<td>Millicell cell culture insert</td>
			<td>Millipore-Sigma</td>
			<td>PICM0RG50</td>
			<td>30 mm, hydrophilic PTFE, 0.4 &#181;m pore, pack of 50.&#160;</td>
		</tr>
		<tr>
			<td>Leica DMi6000 microscope</td>
			<td>Leica Microsystems</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinning disk confocal head Ultraview Vox</td>
			<td>Perkin Elmer</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Volocity 6.0 acquisition software</td>
			<td>Improvision/Perkin Elmer</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>LiveCell Stage top incubation system</td>
			<td>Pathology devices</td>
			<td>LC30030</td>
			<td>Provides Temperature, CO2 and humidity control.&#160;</td>
		</tr>
		<tr>
			<td>SP8 confocal microscope</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>mCherry-Lifeact-7</td>
			<td>Addgene</td>
			<td>54491</td>
			<td>Gift from Michael Davidson</td>
		</tr>
		<tr>
			<td>Fast Green FCF</td>
			<td>Millipore-Sigma</td>
			<td>F7258-25G</td>
			<td>25G bottle, certified by the Biological Stain Commission</td>
		</tr>
	</tbody>
</table>

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.

In this section, we provide representative results obtained following the ex utero electroporation of a control plasmid, or an experimental plasmid targeting a gene of interest, in the MGE of e13.5 mouse embryos followed by organotypic slice cultures incubated at 37 &#176;C for 48 h (for time-lapse imaging) or 72 h (for fixation and immunohistochemical labeling) (see Figure 1B for schematic protocol). Representative examples of INs migrating from an ...

Watch this Scientific Journal Video about Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons at JoVE.com

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.

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 ...

ex-utero-electroporation-organotypic-slice-cultures-embryonic-mouse

Research

JoVE Journal

Neuroscience

9.9K Views.  Centre de recherche du CHU Sainte-Justine. 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.

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

Organotypic Hippocampal Slice Cultures

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the strength of their connections after brief periods of strong activation. This phenomenon, known as long-term potentiation (LTP) can last for hours or days and has become the best candidate mechanism for learning and memory 2. In addition, the well defined anatomy and connectivity of the hippocampus 3 has made it a classical model system to study synaptic transmission and synaptic plasticity4.
As our understanding of the physiology of hippocampal synapses grew and molecular players became identified, a need to manipulate synaptic proteins became imperative. Organotypic hippocampal cultures offer the possibility for easy gene manipulation and precise pharmacological intervention but maintain synaptic organization that is critical to understanding synapse function in a more naturalistic context than routine culture dissociated neurons methods.
Here we present a method to prepare and culture hippocampal slices that can be easily adapted to other brain regions. This method allows easy access to the slices for genetic manipulation using different approaches like viral infection 5,6 or biolistics 7. In addition, slices can be easily recovered for biochemical assays 8, or transferred to microscopes for imaging 9 or electrophysiological experiments 10.

We describe a method to prepare organotypic hippocampal slices that can be easily adapted to other brain regions. Brain slices are laid on porous membranes and culture media is allowed to form an interface. This method preserves the gross architecture of the hippocampus for up to 2 weeks in culture.

The hippocampus, a component of the limbic system, plays important roles in long-term memory and spatial navigation 1. Hippocampal neurons can modify the ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

The mouse is an excellent model organism to study mammalian brain development due to the abundance of molecular and genetic data. However, the developing mouse brain is not suitable for easy manipulation and imaging in vivo since the mouse embryo is inaccessible and opaque. Organotypic slice cultures of embryonic brains are therefore widely used to study murine brain development in vitro. Ex-vivo manipulation or the use of transgenic mice allows the modification of gene expression so that subpopulations of neuronal or glial cells can be labeled with fluorescent proteins. The behavior of labeled cells can then be observed using time-lapse imaging. Time-lapse imaging has been particularly successful for studying cell behaviors that underlie the development of the cerebral cortex at late embryonic stages 1-2. Embryonic organotypic slice culture systems in brain regions outside of the forebrain are less well established. Therefore, the wealth of time-lapse imaging data describing neuronal cell migration is restricted to the forebrain 3,4. It is still not known, whether the principles discovered for the dorsal brain hold true for ventral brain areas. In the ventral brain, neurons are organized in neuronal clusters rather than layers and they often have to undergo complicated migratory trajectories to reach their final position. The ventral midbrain is not only a good model system for ventral brain development, but also contains neuronal populations such as dopaminergic neurons that are relevant in disease processes. While the function and degeneration of dopaminergic neurons has been investigated in great detail in the adult and ageing brain, little is known about the behavior of these neurons during their differentiation and migration phase 5. We describe here the generation of slice cultures from the embryonic day (E) 12.5 mouse ventral midbrain. These slice cultures are potentially suitable for monitoring dopaminergic neuron development over several days in vitro. We highlight the critical steps in generating brain slices at these early stages of embryonic development and discuss the conditions necessary for maintaining normal development of dopaminergic neurons in vitro. We also present results from time lapse imaging experiments. In these experiments, ventral midbrain precursors (including dopaminergic precursors) and their descendants were labeled in a mosaic manner using a Cre/loxP based inducible fate mapping system 6.

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

A method to generate organotypic slices from the E12.5 murine embryonic midbrain is described. The organotypic slice cultures can be used to observe the behavior of dopaminergic neurons or other ventral midbrain neurons.

The  mouse is an excellent model organism to study mammalian brain development due  to the abundance of molecular and genetic data. However, the ...

The Analysis of Purkinje Cell Dendritic Morphology in Organotypic Slice Cultures

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly oriented in the sagittal plane and develops mostly in the postnatal period in small rodents 3. Furthermore, several antibodies are available which selectively and intensively label Purkinje cells including all processes, with anti-Calbindin D28K being the most widely used. For viewing of dendrites in living cells, mice expressing EGFP selectively in Purkinje cells 11 are available through Jackson labs. Organotypic cerebellar slice cultures cells allow easy experimental manipulation of Purkinje cell dendritic development because most of the dendritic expansion of the Purkinje cell dendritic tree is actually taking place during the culture period 4. We present here a short, reliable and easy protocol for viewing and analyzing the dendritic morphology of Purkinje cells grown in organotypic cerebellar slice cultures. For many purposes, a quantitative evaluation of the Purkinje cell dendritic tree is desirable. We focus here on two parameters, dendritic tree size and branch point numbers, which can be rapidly and easily determined from anti-calbindin stained cerebellar slice cultures. These two parameters yield a reliable and sensitive measure of changes of the Purkinje cell dendritic tree. Using the example of treatments with the protein kinase C (PKC) activator PMA and the metabotropic glutamate receptor 1 (mGluR1) we demonstrate how differences in the dendritic development are visualized and quantitatively assessed. The combination of the presence of an extensive dendritic tree, selective and intense immunostaining methods, organotypic slice cultures which cover the period of dendritic growth and a mouse model with Purkinje cell specific EGFP expression make Purkinje cells a powerful model system for revealing the mechanisms of dendritic development.

We present a protocol that permits to view and to quantitatively asses the morphology of the dendritic tree of individual Purkinje cells grown in organotypic cerebellar slice cultures. This protocol is intended to promote studies on the mechanisms of Purkinje cell dendritic development.

Purkinje cells are an attractive model system for studying dendritic development, because they have an impressive dendritic tree which is strictly ...

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for tissue formation during embryonic development and tissue homeostasis during adulthood 1. Currently, great efforts are invested towards controlling the switch of somatic stem cells from expansion to differentiation because this is thought to be fundamental for developing novel strategies for regenerative medicine 1,2. However, a major challenge in the study and use of somatic stem cell is that their expansion has been proven very difficult to control.
Here we describe a system that allows the control of neural stem/progenitor cell (altogether referred to as NSC) expansion in the mouse embryonic cortex or the adult hippocampus by manipulating the expression of the cdk4/cyclinD1 complex, a major regulator of the G1 phase of the cell cycle and somatic stem cell differentiation 3,4. Specifically, two different approaches are described by which the cdk4/cyclinD1 complex is overexpressed in NSC in vivo. By the first approach, overexpression of the cell cycle regulators is obtained by injecting plasmids encoding for cdk4/cyclinD1 in the lumen of the mouse telencephalon followed by in utero electroporation to deliver them to NSC of the lateral cortex, thus, triggering episomal expression of the transgenes 5-8. By the second approach, highly concentrated HIV-derived viruses are stereotaxically injected in the dentate gyrus of the adult mouse hippocampus, thus, triggering constitutive expression of the cell cycle regulators after integration of the viral construct in the genome of infected cells 9. Both approaches, whose basic principles were already described by other video protocols 10-14, were here optimized to i) reduce tissue damage, ii) target wide portions of very specific brain regions, iii) obtain high numbers of manipulated cells within each region, and iv) trigger high expression levels of the transgenes within each cell. Transient overexpression of the transgenes using the two approaches is obtained by different means i.e. by natural dilution of the electroporated plasmids due to cell division or tamoxifen administration in Cre-expressing NSC infected with viruses carrying cdk4/cyclinD1 flanked by loxP sites, respectively 9,15.
These methods provide a very powerful platform to acutely and tissue-specifically manipulate the expression of any gene in the mouse brain. In particular, by manipulating the expression of the cdk4/cyclinD1 complex, our system allows the temporal control of NSC expansion and their switch to differentiation, thus, ultimately increasing the number of neurons generated in the mammalian brain. Our approach may be critically important for basic research and using somatic stem cells for therapy of the mammalian central nervous system while providing a better understanding of i) stem cell contribution to tissue formation during development, ii) tissue homeostasis during adulthood, iii) the role of adult neurogenesis in cognitive functions, and perhaps, iv) better using somatic stem cells in models of neurodegenerative diseases.

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Controlling the expansion of somatic stem cells is a major factor hampering their study and use in therapy. Here we describe a system to temporally control neural stem cells expansion during development and adulthood, which can be used to increase the number of neurons generated in the mouse brain.

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for ...

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 ...

Subtype-selective Electroporation of Cortical Interneurons

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression of genes of interest to specific neuronal subtypes has proven remarkably challenging due to the relative scarcity of specific promoter elements. GABAergic interneurons constitute a neuronal population with extensive genetic and morphological diversity. Indeed, more than 11 different subtypes of GABAergic interneurons have been characterized in the mouse cortex1. Here we present an adapted protocol for selective targeting of GABAergic populations. We achieved subtype selective targeting of GABAergic interneurons by using the enhancer element of the homeobox transcription factors Dlx5 and Dlx6, homologues of the Drosophila distal-less (Dll) gene2,3, to drive the expression of specific genes through in utero electroporation.

This procedure shows how to target interneurons in the developing mouse forebrain by means of in utero electroporation. This technique was particularly efficient to achieve selective gene expression in interneuron subtypes destined to the superficial layers of the cortex.

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression ...

Organotypic Slice Cultures to Study Oligodendrocyte Dynamics and Myelination

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During embryonic and postnatal development they actively proliferate and generate myelinating oligodendrocytes. These cells have commonly been studied in primary dissociated cultures, neuron cocultures, and in fixed tissue. Using newly available transgenic mouse lines slice culture systems can be used to investigate proliferation and differentiation of oligodendrocyte lineage cells in both gray and white matter regions of the forebrain and cerebellum. Slice cultures are prepared from early postnatal mice and are kept in culture for up to 1 month. These slices can be imaged multiple times over the culture period to investigate cellular behavior and interactions. This method allows visualization of NG2 cell division and the steps leading to oligodendrocyte differentiation while enabling detailed analysis of region-dependent NG2 cell and oligodendrocyte functional heterogeneity. This is a powerful technique that can be used to investigate the intrinsic and extrinsic signals influencing these cells over time in a cellular environment that closely resembles that found in vivo.

A technique to study NG2 cells and oligodendrocytes using a slice culture system of the forebrain and cerebellum is described. This method allows examination of the dynamics of proliferation and differentiation of cells within the oligodendrocyte lineage where the extracellular environment can be easily manipulated while maintaining tissue cytoarchitecture.

NG2 expressing cells (polydendrocytes, oligodendrocyte precursor cells) are the fourth major glial cell population in the central nervous system. During ...

The Analysis of Neurovascular Remodeling in Entorhino-hippocampal Organotypic Slice Cultures

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional deficits in the affected patients. Despite intensive research efforts, there is still no effective treatment option available that reduces neuronal injury and protects neurons in the ischemic areas from delayed secondary death. Research in this area typically involves the use of elaborate and problematic animal models. Entorhino-hippocampal organotypic slice cultures challenged with oxygen and glucose deprivation (OGD) are established in&#160;vitro models which mimic cerebral ischemia. The novel aspect of this study is that changes of the brain blood vessels are studied in addition to neuronal changes and the reaction of both the neuronal compartment and the vascular compartment can be compared and correlated. The methods presented in this protocol substantially broaden the potential applications of the organotypic slice culture approach. The induction of OGD or hypoxia alone can be applied by rather simple means in organotypic slice cultures and leads to reliable and reproducible damage in the neural tissue. This is in stark contrast to the complicated and problematic animal experiments inducing stroke and ischemia in vivo. By broadening the analysis to include the study of the reaction of the vasculature could provide new ways on how to preserve and restore brain functions. The slice culture approach presented here might develop into an attractive and important tool for the study of ischemic brain injury and might be useful for testing potential therapeutic measures aimed at neuroprotection.

A protocol for entorhino-hippocampal organotypic slice cultures, which allows reproducing many aspects of ischemic brain injury, is presented. By studying changes of the neurovasculature in addition to changes in the neurons, this protocol is a versatile tool to study plastic changes in neural tissue after injury.

Ischemic brain injury is among the most common and devastating conditions compromising proper brain function and often leads to persisting functional ...

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 ...