This work was funded by a Wellcome Trust Project Grant awarded to P.D. and G.L. (089236/Z/09/Z). S.G. was supported by a Biotechnology and Biological Sciences Research Council PhD studentship. We thank Matthieu Vermeren for the kind gift of the shRNA vector and Jennifer Shieh for valuable advice on neuroblast nucleofection.

This procedure is in accordance with the UK Home Office Regulations (Animal Scientific Procedures Act, 1986). Scientists should follow the guidelines established and approved by their institutional and national animal regulatory organizations.
1. Dissection and Dissociation of Rat RMS Neuroblasts
<ol>
	<li>Prepare the solutions required for RMS dissection and dissociation:</li>
</ol>
Dissection medium (100 ml) 
Hank&#39;s Balanced Salt Solution (HBSS) - 98.5 ml 
5 M HEPES pH 7.4 - 0.5 ml 
Penicillin-streptomycin (10,000 units/ml and 10,000 &#181;g/ml) - 1 ml
Dissociation medium (2 ml) 
HBSS - 1.760 ml 
10x Trypsin (2.5%) - 200 &#181;l 
DNAse1 (1 mg/ml) - 40 &#181;l
Dulbecco Modified Eagle&#39;s Medium (DMEM) + 10% Fetal Calf Serum (FCS) (40 ml) 
DMEM - 36 ml 
FCS - 4 ml
Complete medium (12 ml)
Neurobasal medium - 11.46 ml 
B27 Supplement - 250 &#181;l 
L-Glutamine (200 mM) - 125 &#181;l 
Glucose (45%) - 165 &#181;l
2. Filter-sterilize the DMEM + 10% FCS and the complete medium and preequilibrate them in a 37 &#176;C /5% CO2 incubator.
<ol>
	<li>Dissection
	<ol>
		<li>Sacrifice a P6-P7 rat litter (about 12 pups) by cervical dislocation and decapitate with scissors.</li>
		<li>Make an anteroposterior incision in the skin along the mid-sagittal suture from the nose to the cerebellum with a scalpel blade. Peel the skin off and repeat the same incision along the skull.</li>
		<li>Gently remove the cranial flaps with forceps and carefully remove the brain with a spatula, taking care to include the olfactory bulbs.</li>
		<li>Cut the most caudal third of the brain and discard it.</li>
		<li>Chop the brain tissue into 1.4 mm&#160;thick coronal slices using a tissue chopper.</li>
		<li>Place slices in dishes containing cold dissection medium and carefully separate them using a needle.</li>
		<li>The RMS appears as a triangular, translucent area in the centre of OB sections and as a small, circular area in more caudal brain slices. Cut the RMS out of each slice with a microsurgical knife, taking care to avoid including surrounding tissue. In P7 rat pups, usually the ~8 most rostral slices (including OB) contain the RMS.</li>
		<li>Collect the RMS fragments with a plastic Pasteur pipette and place them in a small dish containing cold dissection medium on ice.</li>
		<li>When the dissection is complete, transfer the RMS fragments into a 15 ml tube with a plastic pipette. Leave fragments to settle at the bottom of the tube.</li>
	</ol>
	</li>
	<li>Dissociation
	<ol>
		<li>Replace the dissection medium with 2 ml of dissociation medium.</li>
		<li>Triturate the RMS fragments by gently pipetting the fragment suspension up and down about 10x using a P1000 pipette.</li>
		<li>Leave the tube with the tissue fragments in a 37 &#176;C water bath for 2 min.</li>
		<li>Pipette the solution again 10x and ensure that fragments have dissociated (the suspension should become cloudy).</li>
		<li>Inactivate the trypsin by adding 5 ml of prewarmed DMEM + 10% FCS.</li>
		<li>Centrifuge the cell suspension at 433 x g for 5 min.</li>
		<li>In the meantime aliquot the required amount of siRNA/DNA into Eppendorf tubes (usually 3-5 &#181;g DNA/shRNA or 5-9 &#181;g siRNA oligo per nucleofection, however the amount of DNA/siRNA may require optimization).</li>
		<li>Remove excess medium and resuspend the cell pellet by gentle pipetting in 5 ml of prewarmed DMEM + 10% FCS.</li>
		<li>Perform a cell count. Expect ~1 x 106 cells per rat pup brain. A minimum of 2.5 x 106 cells are required for each nucleofection, while optimal results are achieved using 3-4 x 106 cells per nucleofection.</li>
		<li>Centrifuge the cell suspension at 433 x g for 5 min. Make sure to remove as much medium as possible.</li>
	</ol>
	</li>
</ol>
2. Nucleofection
<ol>
	<li>Immediately resuspend the cell pellet in rat (or mouse if using mouse cells) neuron nucleofection solution previously incubated at room temperature. Use 100 &#181;l per nucleofection. Note: typically a rat litter of 12 pups is sufficient to perform 4 nucleofections and a mouse litter (12 pups) is sufficient to perform 2 nucleofections.</li>
	<li>Transfer 100 &#181;l of the cell suspension to each Eppendorf tube containing siRNA/DNA and mix gently 2-3x by pipetting with a P200 pipette.</li>
	<li>Add the sample (cell-DNA/siRNA suspension) to the bottom of the nucleofectioncuvette, taking care to avoid bubbles.</li>
	<li>Nucleofect using program G-013 (for rat cells) or O-005 (for mouse cells). One nucleofection takes approximately 5 sec.</li>
	<li>Quickly add 1 ml of prewarmed DMEM + 10% FCS to the nucleofected sample.</li>
	<li>Repeat steps 2.4 and 2.5 for all other samples. Note: for optimal results, the entire nucleofection procedure should last no longer than 5 min.</li>
	<li>Transfer each sample to a 15 ml tube containing 5 ml of prewarmed DMEM + 10% FCS using the plastic pipette provided by the nucleofection kit. Avoid transferring any cellular debris to the tube.</li>
	<li>Centrifuge samples at 433 x g for 5 min.</li>
	<li>Carefully remove all excess medium and resuspend the pellet in 25-30 &#181;l of prewarmed DMEM + 10% FCS using a P20 pipette. Do not use more than 30 &#181;l of medium.</li>
	<li>Pipette the suspension as a drop onto the inner side of a p35 dish lid.</li>
	<li>Invert the lid over the p35 dish containing 2 ml of complete medium (see also Figure 1).</li>
	<li>Leave in the incubator (37 &#176;C/5% CO2) for at least 5 hr and up to 7 hr. Longer incubation time allows better reaggregation of cell clusters.</li>
	<li>Transfer the hanging drops from the lid into the complete medium in the dish using a P1000 pipette with a cut tip.</li>
	<li>Incubate at 37 &#176;C/5% CO2 for 24 hr for DNA nucleofections and 48 hr for siRNA/ shRNA nucleofections.</li>
</ol>
3. Embedding
<ol>
	<li>Prepare complete medium (25 ml) and preequilibrate it at 37 &#176;C/5% CO2 for a few hours.</li>
	<li>Take out the frozen aliquots of basement membrane matrix from -80 &#176;C freezer and thaw on ice in the cold room.</li>
	<li>For each nucleofection prepare a 6 cm dish containing up to eight 13 mm sterile coverlips.</li>
	<li>Place the dishes on an ice box covered with cling film. It is important to keep the coverslips cool to prevent matrix solidification during the embedding procedure.</li>
	<li>To maintain humidity, place a strip of damp tissue inside a 15 cm dish that will be used to hold up to three 6 cm dishes containing the embedded neuroblasts.</li>
	<li>Add complete medium to the thawed matrix in a 1:3 ratio. For example, mix 40 &#181;l of complete medium with 120 &#181;l of matrix by pipetting. This amount of matrix is sufficient for embedding aggregates on eight 12 mm coverslips.</li>
	<li>Transfer the reaggregated cell clusters to a 15 ml tube and centrifuge at 433 x g for 5 min.</li>
	<li>Remove excess medium and resuspend the pellet in 10 &#181;l of complete medium.</li>
	<li>Place 2 &#181;l of cell aggregate suspension onto each sterile coverslip and add 18 &#181;l of matrix/complete medium mixture. Use the pipette tip to spread the matrix over the entire coverslip.</li>
	<li>Immediately place the 6 cm dish containing the coverslips in the 15 cm dish and leave in the incubator (37&#176;C /5% CO2) for 15-20 min. When the matrix has solidified, gently add 5 ml complete medium to each 6 cm dish taking care to push down any floating coverslip with a pipette tip.</li>
	<li>Incubate for 24 hr at 37 &#176;C /5% CO2 to let neuroblasts migrate out of the cell aggregates.</li>
</ol>
4. 3D Migration Assay
<ol>
	<li>Prepare the solutions required for immunostaining.
	<ol>
		<li>Prepare the block solution: 
		Goat block solution (50 ml) 
		Phosphate Buffered Saline (PBS) - 5 ml 
		Goat serum - 7.5 ml 
		10% Triton X-100 - 1.5 ml 
		BSA - 50 mg 
		H2O - 36 ml</li>
		<li>Filter and store at 4 &#176;C.</li>
		<li>Prepare the fixing solution: 
		Fixing solution (100 ml) 
		Paraformaldehyde (PFA) - 4 g CAUTION: always handle PFA under a hood 
		Sucrose - 20 g (optional) 
		PBS up to 100 ml</li>
		<li>On a hot plate and under constant stirring dissolve the PFA in 80 ml PBS maintained at 65 &#176;C.</li>
		<li>Once the PFA has dissolved, add 20 g of sucrose.</li>
		<li>Adjust pH to 7.4 (usually by adding ~60 &#181;l 1 M NaOH per 100 ml of solution).</li>
		<li>Bring up to a total volume of 100 ml with PBS.</li>
	</ol>
	</li>
	<li>Immunostaining
	<ol>
		<li>Place coverslips in a 24-well plate.</li>
		<li>Rinse coverslips with PBS 2x.</li>
		<li>Fix the RMS neuroblast aggregates with fixing solution for 45 min at room temperature.</li>
		<li>Rinse coverslips with PBS 3x (5 min/wash - on a rocking platform).</li>
		<li>Block for 30-60 min&#160;with goat block solution.</li>
		<li>Dilute primary antibodies in goat block solution and incubate overnight at 4 &#176;C. (If desired, fluorescent phalloidin (1:400) and Hoechst dye (1:10,000) can also be added to the primary antibody solution to visualize filamentous actin and nuclei).</li>
		<li>Rinse coverslips with PBS 3x&#160;(5 min/wash).</li>
		<li>Dilute secondary antibodies in goat block solution and incubate for 2 hr at room temperature.</li>
		<li>Rinse coverslips with PBS 3x&#160;(5 min/wash)</li>
		<li>Mount coverslips with fluorescent mounting medium and leave to dry overnight at room temperature.</li>
	</ol>
	</li>
	<li>Migration Analysis
	<ol>
		<li>Capture images of fixed RMS neuroblast aggregates with a fluorescent microscope using a 10X&#160;objective. Include a scale bar in a sample image.</li>
		<li>To set up the scale for quantification, measure the scale bar in the image by selecting the &#39;Straight line&#39; tool on the ImageJ tool bar.</li>
		<li>Choose the &#39;Analyze&#39; option and click on &#39;set the scale&#39;.</li>
		<li>In the scale window set the &#39;known distance&#39; and tick the &#39;global&#39; box to keep the same settings for all measurements.</li>
		<li>Use the &#39;Segmented line&#39; tool on the ImageJ tool bar to measure the distance from the edge of the aggregate to the furthest migrated neuroblast in 6 different sectors around the entire aggregate (Figure 3B). Consider only isolated aggregates for analysis.</li>
		<li>Calculate an average migration distance from the 6 values obtained for each aggregate.</li>
		<li>Measure 10-20 aggregates for each condition in every independent experiment and pool results from a minimum of three independent experiments. Always include a nucleofection control (e.g.&#160;GFP or a control sh/siRNA).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The migration of neuroblasts along the RMS to the final location in the OB is a fundamental step in postnatal neurogenesis. However, the molecular mechanisms controlling this complex process are far from being fully understood.
The experimental procedure described here allows the study of neuroblast migration in vitro. We have adapted a previously published protocol for isolating RMS neuroblasts from early postnatal mouse or rat25. To achieve optimal results it is important to master the dissection step, since it is crucial to keep the time interval between dissection and nucleofection to a minimum. After nucleofection, neuroblasts can be reaggregated, embedded in a three-dimensional matrix and left to migrate over a 24 hr period. Alternatively, for purposes other than migration (e.g. immunofluorescence or western blot analysis), cells can be immediately plated after nucleofection on polyornithine/laminin-coated coverslips, where they survive up to 4-5 days. Mouse and rat neuroblasts migrate in Matrigel to a similar extent, however mouse cells appear to have a stronger tendency to migrate in chains than rat cells.
Depending on the aim of the study, neuroblasts can be nucleofected with different plasmids encoding fluorescent proteins or wild type/mutant proteins of interest. For optimal protein expression plasmids with a CAG promoter (&#946;-actin promoter with CMV enhancer and &#946;-globin poly-A tail)26 are highly recommended. Moreover, siRNA oligos or shRNA plasmids can be nucleofected to knockdown targets of interest. Effective protein depletion can be visualized by immunofluorescence or by western blot (usually lysing embedded aggregates from 1 rat pup with 50 &#181;l of standard lysis buffer).
Nucleofection is a relatively simple method to transfect primary neuroblasts, offers an easier and faster alternative to viral vector-mediated transfection, and can achieve high (~70-80%) transfection efficiency. It is critical to work quickly during the nucleofection procedure, since leaving neuroblasts in the nucleofection solution for a prolonged time drastically reduces cell viability.
The average cell yield from RMS dissection is relatively low for P7 mice (~5 x 105 cells/brain) in comparison to P7 rats (~1 x 106 cells/brain) and at least 3 x 106 cells per nucleofection are required to achieve transfection with ~50% efficiency. Moreover, rat neuroblasts appear to resist better to nucleofection compared to mouse neuroblasts. Therefore, early postnatal (P6-P7) rat pups might represent a convenient neuroblast source, also considering that the organization of rat and mouse RMS are remarkably similar27 and that the extent of rat and mouse neuroblast migration in vitro is also comparable. It is advisable not to keep the reaggregated clusters of nucleofected neuroblasts in suspension for longer than 48 hr to avoid abnormal effects on cell morphology and migration (our unpublished observations).
The 3D assay described here can be used to quantify neuroblast migration at a fixed time point after embedding in matrix (e.g. 24 hr). Aggregates of different sizes can be used in the analysis, since there is no significant correlation between the size of aggregates and migration distance (our unpublished observations). To visualize and further investigate the dynamics of neuroblast migration, time-lapse imaging can be used. It is recommended to carry out the migration analysis within a 24 hr interval after embedding, since the speed of neuroblasts appears to drastically decrease at longer time points (our unpublished observations).
There are some limitations to this protocol. First, nucleofection can so far be used for early postnatal rodent neuroblasts, while infection with viral vectors remains the most efficient transfection method for adult neuroblasts28. Second, the in vitro migration assay does not fully reproduce the complex architecture of the RMS observed in vivo. Indeed, although neuroblasts maintain the ability to migrate in a similar way to their in vivo counterparts, in the experimental setup described here they lack interactions with other RMS components such as astrocytes and blood vessels, which also contribute to regulate their motility9,29,30. This issue may be addressed in the future by optimization of three-dimensional coculture model systems.
In conclusion, combining nucleofection with a 3D migration assay represents a valuable tool to better understand the molecular mechanisms underlying neuroblast migration. This experimental procedure provides an initial, fast and relatively simple method to evaluate the role of candidate regulators of neuroblast migration, which can be further validated by other approaches like in vivo postnatal electroporation and time-lapse imaging of brain slice cultures28,31,32.

In the postnatal mammalian brain, generation of new neurons (neurogenesis) occurs throughout life and is restricted to two neurogenic niches: the subventricular zone (SVZ) of the lateral ventricles and the subgranular zone of the dentate gyrus of the hippocampus1. Several recent studies have shown the important role of adult neurogenesis in facilitating learning and memory tasks2,3. Moreover, evidence of proliferation and recruitment of neural progenitors following brain injury4-7 raises the possibility of pharmacological activation of neurogenesis in neural repair.
Postnatal neurogenesis is strictly regulated in all its phases, which include neural progenitor proliferation, migration, differentiation, survival, and final synaptic integration of newly born neurons8. Neural progenitors (neuroblasts) derived from stem cells in the SVZ migrate over a great distance through the rostral migratory stream (RMS) towards the olfactory bulb (OB) where they mature into functional neurons9. Migratory neuroblasts are predominantly unipolar, with an elongated cell body extending a single leading process. These cells move in chains in a collective manner, sliding over one another10. Migration is a crucial step for the subsequent maturation of SVZ-derived progenitors into functional neurons11 and is controlled by multiple factors and guidance molecules including: polysialylated neural cell adhesion molecule (PSA-NCAM)12, Ephrins13, integrins14, Slits15, growth factors16 and neurotransmitters17, however the molecular mechanisms underlying this process are not fully understood. Investigating the intracellular signaling pathways regulating neuroblast migration will not only provide a better understanding of adult neurogenesis, but will also contribute to the development of the new therapeutic approaches to promote brain repair.
This manuscript describes a detailed protocol to study the role of candidate regulators of neuroblast migration in vitro using nucleofection and a 3D migration assay. Nucleofection is a cell transfection technique based on an improved method of electroporation. Cell-type specific electrical current and nucleofection solution allow the transfer of polyanionic macromolecules such as DNA and shRNA vectors and siRNA oligonucleotides directly into the cell nucleus and permit transfection of slowly dividing or mitotically inactive cells like embryonic and mammalian neurons18. This method is fast, relatively easy to perform and results in highly reproducible transfection of a broad range of cell types including primary neuroblasts and neurons19-21.
Dissociation of RMS tissue allows the isolation of migratory neuroblasts, which can be successfully nucleofected with DNA/shRNA vectors or siRNA oligos targeting genes of interest. Following nucleofection, neuroblasts are reaggregated in hanging drops and subsequently embedded in a three-dimensional Matrigel matrix. These conditions allow neuroblasts to migrate out of the cell aggregates recapitulating the migration mode observed in vivo, thus providing an excellent model system to investigate signaling pathways involved in neuroblast migration and to assess the influence of pharmacological treatments on the motility of these cells.

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			<td height="21" style="height:21px;width:688px;">2.5% Trypsin-EDTA (10x)</td>
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			<td>store 200 &#181;l aliquots at -20 &#176;C</td>
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			<td style="width:255px;">LK003170</td>
			<td>&#8805;1,000 units per vial; store 50 &#181;l aliquots at -20 &#176;C</td>
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			<td style="width:255px;">G8769</td>
			<td></td>
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			<td height="21" style="height:21px;">Matrigel Basement Membrane Matrix, Growth Factor Reduced (GFR), Phenol Red-free, 10 ml, LDEV-Free</td>
			<td style="width:339px;">BD Biosciences</td>
			<td>356231</td>
			<td>prepare 120 &#181;l aliquots at 4 &#176;C, then store at -80 &#176;C</td>
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			<td height="21" style="height:21px;width:688px;">PFA</td>
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			<td style="width:339px;">Sigma-Aldrich</td>
			<td style="width:255px;">69023</td>
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			<td height="21" style="height:21px;width:688px;">Triton X-100</td>
			<td style="width:339px;">VWR International Ltd.</td>
			<td style="width:255px;">306324N</td>
			<td></td>
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			<td height="21" style="height:21px;width:688px;">BSA</td>
			<td style="width:339px;">Fisher Chemical</td>
			<td style="width:255px;">BPE9701-100</td>
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			<td height="21" style="height:21px;width:688px;">Dako fluorescence mounting medium</td>
			<td style="width:339px;">Dako</td>
			<td style="width:255px;">S3023</td>
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			<td height="21" style="height:21px;width:688px;">Rat neuron nucleofection kit</td>
			<td style="width:339px;">Lonza</td>
			<td style="width:255px;">VPG-1003</td>
			<td></td>
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			<td height="21" style="height:21px;width:688px;">Mouse neuron nucleofection kit</td>
			<td style="width:339px;">Lonza</td>
			<td style="width:255px;">VPG-1001</td>
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			<td height="21" style="height:21px;width:688px;">Microsurgical knife</td>
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			<td style="width:255px;">7516</td>
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			<td height="21" style="height:21px;width:688px;">McIlwain tissue chopper</td>
			<td style="width:339px;">The Mickle Laboratory Engineering Company</td>
			<td style="width:255px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;width:688px;">Nucleofector II</td>
			<td style="width:339px;">Lonza</td>
			<td style="width:255px;"></td>
			<td></td>
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nucleofection of rodent neuroblasts to study neuroblast migration in vitro

The subventricular zone (SVZ) located in the lateral wall of the lateral ventricles plays a fundamental role in adult neurogenesis. In this restricted area of the brain, neural stem cells proliferate and constantly generate neuroblasts that migrate tangentially in chains along the rostral migratory stream (RMS) to reach the olfactory bulb (OB). Once in the OB, neuroblasts switch to radial migration and then differentiate into mature neurons able to incorporate into the preexisting neuronal network. Proper neuroblast migration is a fundamental step in neurogenesis, ensuring the correct functional maturation of newborn neurons. Given the ability of SVZ-derived neuroblasts to target injured areas in the brain, investigating the intracellular mechanisms underlying their motility will not only enhance the understanding of neurogenesis but may also promote the development of neuroregenerative strategies.
This manuscript describes a detailed protocol for the transfection of primary rodent RMS postnatal neuroblasts and the analysis of their motility using a 3D in vitro migration assay recapitulating their mode of migration observed in vivo. Both rat and mouse neuroblasts can be quickly and efficiently transfected via nucleofection with either plasmid DNA, small hairpin (sh)RNA or short interfering (si)RNA oligos targeting genes of interest. To analyze migration, nucleofected cells are reaggregated in &#39;hanging drops&#39; and subsequently embedded in a three-dimensional matrix. Nucleofection per se does not significantly impair the migration of neuroblasts. Pharmacological treatment of nucleofected and reaggregated neuroblasts can also be performed to study the role of signaling pathways involved in neuroblast migration.

Neuroblasts can be successfully isolated from dissected RMS tissue (Figure 1A) and embedded in a three-dimensional matrix. Cells isolated from either rat or mouse postnatal RMS are immunopositive for migratory neuroblast markers, such as doublecortin (DCX), &#946;III tubulin or PSA-NCAM (Figures 1B-C).&#160;
Dissociated neuroblasts can be efficiently nucleofected with DNA (e.g. a GFP-encoding plasmid, Figure 2) or shRNA plasmids (Figure 4) to achieve protein depletion, which can be assessed by western blot analysis (Figure 4B) or immunofluorescence (not shown).
Cells nucleofected with GFP-encoding plasmids migrate radially out reaggregated neuroblast clusters (Figure 3A). Quantification of relative migrated distance 24 hr post embedding (Figure 3B) shows no difference in migration between GFP-positive cells and GFP-negative, nonnucleofected cells (Figure 3C), indicating that nucleofection per se does not disrupt migration. There is also no significant difference in the extent of migration between nucleofected neuroblasts and neuroblasts directly migrating out of RMS explants (data not shown).
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50989/50989fig1.jpg" width="500px" /> 
Figure 1. Dissection of RMS neuroblasts. (A) Schematic representation of RMS neuroblast dissection. For detailed description please refer to the text. (B) Isolated rat RMS cells are immunopositive for the migratory neuroblast makers DCX and &#946;lll tubulin. Bar, 20 &#181;m. (C) Cells migrating out of mouse RMS explants express the migratory neuroblast markers DCX and PSA-NCAM. Bar, 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50989/50989fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="6in" src="/files/ftp_upload/50989/50989fig2.jpg" width="500px" /> 
Figure 2. Mouse neuroblast nucleofection. Dissociated mouse RMS neuroblasts were nucleofected with pMAX-GFP, reaggregated, embedded in a three-dimensional matrix and allowed to migrate for 6 hr. Neuroblasts migrating out of a reaggregated cell cluster (top, phase contrast pictures) show high transfection efficiency (bottom, GFP channel pictures). The right column panels show higher magnification pictures corresponding to the insets highlighted in the left column panels. Bars, 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/50989/50989fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50989/50989fig3.jpg" width="500px" /> 
Figure 3. 3D Migration assay. (A) Rat neuroblasts were nucleofected with pMAX-GFP (GFP) or pCAG-IRES-EGFP22 (EV), reaggregated, embedded in matrix and left to migrate for 24 hr. Cells were then fixed and immunostained for GFP (green) and &#946;III tubulin (red). Bar, 50 &#181;m. (B) Measuring migration distance using ImageJ. The reaggregated cell cluster is divided into 6 equal sectors. The distance between the edge of the cluster (dotted line) and the furthest migrated cell is measured for each sector. (C) Quantification of the relative distance migrated by nucleofected cells (GFP-positive) and control, nonnucleofected cells (GFP-negative). <a href="https://www.jove.com/files/ftp_upload/50989/50989fig3highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50989/50989fig4.jpg" width="500px" /> 
Figure 4. Monitoring neuroblast migration after shRNA nucleofection. (A) Rat neuroblasts were nucleofected with a control shRNA vector (pCA-b-EGFPm5 Silencer 3, which also expresses EGFP23) or the same vector containing a shRNA targeting fascin, an actin-bundling protein24. Cells were reaggregated over 48 hr, embedded in matrix and left to migrate for 24 hr. Aggregates were then fixed and immunostained for GFP (green) and &#946;III tubulin (red). Bar, 50 &#181;m. (B) Effective fascin depletion can be detected 50 hr after shRNA nucleofection by western blot analysis. Actin is shown here as a loading control. (C) Quantitative analysis of relative migration distance showing that fascin depletion significantly impairs neuroblast migration (mean &#177; SEM; **p&#60;0.01; n=3 independent experiments). <a href="https://www.jove.com/files/ftp_upload/50989/50989fig4highres.jpg" target="_blank">Click here to view larger image</a>.

Watch this Scientific Journal Video about Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro at JoVE.com

Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro

Neuroblast migration is a crucial step in postnatal neurogenesis. The protocol described here can be used to investigate the role of candidate regulators of neuroblast migration by employing DNA/small hairpin RNA (shRNA) nucleofection and a 3D migration assay with neuroblasts isolated from the rodent postnatal rostral migratory stream.

Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro

The subventricular zone (SVZ) located in the lateral wall of the lateral ventricles plays a fundamental role in adult neurogenesis. In this restricted ...

nucleofection-rodent-neuroblasts-to-study-neuroblast-migration

Research

JoVE Journal

Neuroscience

11.4K Görüntüleme.  King's College London.  Neuroblast göç postnatal nörojenezindeki çok önemli bir adımdır. Protokol Burada açıklanan DNA / küçük firkete RNA (ShRNA) nucleofection ve kemirgen doğum sonrası rostral göçmen akımından izole nöroblast ile 3D göç deneyi kullanılarak neuroblast göç aday düzenleyicilerinin rolünü araştırmak için kullanılabilir. 

Video: Nucleofection of Rodent Neuroblasts to Study Neuroblast Migration In vitro

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

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 ...

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 ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

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 ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering the molecular signals that cooperatively guide neuronal migration in the embryonic brain is therefore important to understand how the complex neural networks form which later support postnatal life. Facial branchiomotor (FBM) neurons in the mouse embryo hindbrain migrate from rhombomere (r) 4 caudally to form the paired facial nuclei in the r6-derived region of the hindbrain. Here we provide a detailed protocol for wholemount ex vivo culture of mouse embryo hindbrains suitable to investigate the signaling pathways that regulate FBM migration. In this method, hindbrains of E11.5 mouse embryos are dissected and cultured in an open book preparation on cell culture inserts for 24 hr. During this time, FBM neurons migrate caudally towards r6 and can be exposed to function-blocking antibodies and small molecules in the culture media or heparin beads loaded with recombinant proteins to examine roles for signaling pathways implicated in guiding neuronal migration.

Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Embryonic neurons are born in the ventricular zone of the neural tube, but migrate to reach appropriate targets. Facial branchiomotor (FBM) neurons are a useful model to study neuronal migration. This protocol describes the wholemount ex vivo culture of mouse embryo hindbrains to investigate mechanisms that regulate FBM migration.

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate targets. Deciphering ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...