MT is supported by a PhD studentship [ref. 092839/Z/10/Z] and CR by a New Investigator Award [ref. 095623/Z/11/Z] from the Wellcome Trust.

1. Optional: Prepare Affi-gel Heparin Beads (Gel Beads) for FBM Attraction Assay
NOTE: Prepare gel beads at least 1 day before starting the explant procedure.
<ol>
	<li>Wash 100 &#956;l gel heparin bead suspension with sterile PBS for 20 min on a roller at room temperature (RT).</li>
	<li>Pellet beads in a table top centrifuge for 5 min at 13,000 x g. Add sterile PBS and repeat the washing procedure 4x.</li>
	<li>After the final wash, remove PBS and soak the beads in a small volume of a sterile solution containing the recombinant protein of choice, taking care to cover the beads with the solution. This protocol uses 100 ng/&#956;l recombinant human VEGF165 in PBS to reproduce a previously published experiment16.</li>
	<li>Incubate the gel heparin beads with the recombinant protein solution for a minimum of 12 hr and a maximum of 1 week on a roller at 4 &#176;C.</li>
</ol>
2. Coating of Culture Inserts
Hindbrain explants are cultured on Corning culture inserts with an 8 &#956;m pore size, or equivalent inserts. Culture inserts can be reused after completion of the protocol, provided they are washed with distilled water, sterilized with ethanol, and stored in 70% ethanol until needed. 
NOTE: The following steps should be carried out in a flow hood under sterile conditions.
<ol>
	<li>Prepare explant culture media consisting of Neurobasal medium supplemented with B27 (20 &#956;l/ml), glucose (6 mg/ml) and penicillin/streptomycin (5 &#956;g/&#956;l).</li>
	<li>Wash culture inserts with sterile PBS for 5 min and dry for 5-10 min under the flow hood.</li>
	<li>Place one culture insert into each individual well of a 12well plate. Note: inserts may need a small push to fit tightly into the well.</li>
	<li>Cover the culture inserts with 10-20 &#956;g/ml mouse laminin in Neurobasal medium and place them in a tissue culture incubator (37 &#176;C, 5% CO2) for 1 hr. Note: Coating should be carried out on the day of explanting.</li>
</ol>
3. Dissection of Hindbrains from E11.5 Mouse Embryos
<ol>
	<li>Cull timed pregnant female mouse with an ethically approved procedure on embryonic day (E) 11.5 and place the uterus containing the embryos in a 100 mm plastic dish with ice-cold L15 medium. 
	NOTE: All dissection steps must be performed in ice-cold L15.</li>
	<li>Using a dissecting microscope and Dumont watchmaker forceps number 5, tear the uterine muscle wall to expose the embryos, release each embryo, sever the umbilical cord, and carefully remove the yolk sac.</li>
	<li>Using a plastic Pasteur pipette with a wide-bore opening, transfer each embryo into a clean plastic dish with ice-cold L15.</li>
	<li>Using Dumont forceps, decapitate the embryo just above the forelimbs. If the experiment requires genotyping of the embryos, collect tissue samples for genomic DNA isolation (e.g. a small piece of yolk sac or tail tip16,20).</li>
	<li>Turn the head dorsal side up and identify the 4th ventricle, which is covered by a thin tissue layer (Figure 2B). Carefully pierce the roofplate and begin to peel it away caudally along the midline over the posterior hindbrain and spinal cord, and rostrally over the midbrain. The hindbrain should now be exposed (Figure 2C).</li>
	<li>Carefully tease away the remaining head mesenchyme and any meninges that are attached to the pial side of the hindbrain (Figure 2D).</li>
	<li>Remove midbrain and spinal cord tissue so that the hindbrain unfurls and can lie flat in an open book preparation (Figure 2E).</li>
	<li>Using a wide-bore plastic Pasteur pipette, transfer each dissected hindbrain to a 12-well plate containing ice-cold L15 and store them on ice until all hindbrains have been dissected.</li>
	<li>Using a wide-bore plastic Pasteur pipette, transfer a single hindbrain to an empty dish, keeping it an open book preparation, ventricular side up, in a droplet of approximate 100 &#956;l L15.</li>
	<li>Transfer a few microliters of the incubated gel heparin beads to the same droplet. Note: Because the sizes of beads are variable, it is advisable to transfer around 10 beads and then choose an optimal size for transplantation into the hindbrain.</li>
	<li>Make a small tear in the hindbrain tissue and carefully insert 1-3 gel beads into the hindbrain tissue at the level of r5/6, about half way between the midline and the lateral edge of the hindbrain, lowering them into the tissue so that the bead is positioned just beneath the hindbrain surface. 
	NOTE: The dissection procedure may take between 5-20 min/hindbrain, depending on experience, and may stretch over an extended period if litters are large; in any case, hindbrains should be in culture no longer than 3 hr post-mortem for good results.</li>
</ol>
4. Hindbrain Explant Culture
<ol>
	<li>Remove the plate containing the culture inserts from the incubator, aspirate the laminin coating solution.</li>
	<li>Place one culture insert into a separate culture dish filled with ice-cold L15 and, using a wide-bore plastic Pasteur pipette, transfer each hindbrain ventral side up onto the culture insert (Figure 2G). The hindbrain should lie completely flat on the insert membrane.</li>
	<li>Carefully lift the culture insert from the dish and dab it several times on clean tissue paper to remove excess liquid. This process ensures that the hindbrain adheres to the culture insert in a flat, open book preparation. If the hindbrain curls up, its tissue may irreversibly grow together.</li>
	<li>Fill the original 12-well plate with 500 &#956;l prewarmed culture media and place the insert back into this well. Carefully adjust the volume with another 400-600 &#956;l of media to just cover the hindbrain, ensuring that the hindbrain does not float off the membrane. If it floats, return to step 4.4 and repeat until the hindbrain remains attached to the membrane.</li>
	<li>At this stage, it is possible to add biological inhibitors of interest to the media to study their effect on the migration of FBM neurons. 
	NOTE: If implanting beads or administering other treatments, it is recommended to maintain at least 2 control explants under normal growth conditions per experiment to ensure that the experiment has been set up successfully.</li>
	<li>Incubate explants for 24-30 hr in a tissue culture incubator (37 &#176;C, 5% CO2).</li>
</ol>
5. Wholemount Immunofluorescent Staining of Hindbrain Explants
<ol>
	<li>Aspirate the media from each well, rinse in PBS and fix for 2 hr at 4 &#176;C with gentle agitation in ice-cold 4% formaldehyde (freshly prepared or freshly thawed 4% paraformaldehyde dissolved in PBS). Note: Do not try to remove hindbrain from the culture insert before fixation is complete.</li>
	<li>Rinse 3x with PBS. Carefully peel the hindbrains from the culture inserts using Dumont forceps. Some explants are difficult to peel off, but can usually be lifted by applying gentle pressure through repeatedly expelling PBS from a pipette.</li>
	<li>Transfer the hindbrains to 2.0 ml round-bottomed tubes for immunofluorescence labelling. Permeabilize hindbrains for 30 min at room temperature (RT) in PBS containing 0.1% TritonX-100 (PBT) with gentle rolling.</li>
	<li>Incubate for 1 hr at RT in PBT containing 10% heat-inactivated normal goat serum with gentle rolling.</li>
	<li>Incubate explants with gentle rolling at 4 &#176;C for 5 days with primary antibody specific for ISL1, diluted 1:100 in PBT containing 1% heat-inactivated normal goat serum.</li>
	<li>Wash the explants at RT 4x with PBT for 15 min each.</li>
	<li>Incubate explants with gentle rolling at RT for 3 hr with fluorophore-conjugated goat anti-mouse antibody (e.g. Alexa Fluor 488 goat anti mouse, diluted 1:200) in PBT containing 1% heat-inactivated normal goat serum.</li>
	<li>Wash the explants 4x at RT with PBT for 15 min each with gentle rolling.</li>
	<li>Postfix the explants in 4% formaldehyde for 30 min at RT.</li>
	<li>Cover a glass slide with 3 layers of black electrical tape and excise with a scalpel a small square of the layered tape to create a pocket for the explants; alternatively, use a depression glass slide.</li>
	<li>Mount each hindbrain in SlowFade reagent into one pocket and cover with a glass coverslip, carefully avoiding to trap air bubbles and image using a laser scanning confocal microscope. 
	NOTE: As an alternative to immunostaining, in situ hybridization with riboprobes that recognize FBMs (i.e. Isl1 or Phox2b) may be used to visualize FBM neurons16,21.</li>
</ol>
Summary of steps and timing 
Timed mating to obtain E11.25 pregnancies: ~14 days 
Optional: Bead preparation (Protocol 1): ~2 hr, on the day before embryo isolation 
Prepare culture inserts and media (Protocol 2): ~30 min, before embryo isolation 
Embryo isolation and hindbrain dissection (steps 3.1-3.4): ~10 min/embryo 
Hindbrain dissection (steps 3.5-3.7): ~5-10 min/hindbrain 
Explant procedure (steps 3.8-3.9): ~5-10 min/hindbrain 
Optional: bead implantation (steps 3.10-3.11): ~5-10 min/hindbrain 
Explant culture (step 4.7): 24 hr 
Fixation for antibody staining (step&#160;5.1): 2 hr 
Staining procedure and imaging (Protocol 5): 5 days

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None of the authors have competing interests or conflicting interests.

This protocol describes the wholemount culture of E11.5 mouse hindbrains in a transwell system to study the migration of FBM neurons. This protocol allows mouse hindbrain motorneurons to be kept alive and migrating for a period of 24 hr, enabling ex vivo manipulation. This method has numerous experimental advantages for investigators seeking to identify the molecular and cellular mechanisms of neuronal migration. Whereas traditional migration assays explant small neural tissue pieces into matrix on culture dishes and enable observation of individual neurons as they respond to exogenous stimuli, a major advantage of the transwell assay is its suitability to manipulate migrating neurons within the host organ environment and therefore a more physiological context. Importantly, substances can be readily applied to the ex vivo hindbrain explants to test their effect on neuronal migration, circumventing possible side effects associated with administering these substances to a pregnant mouse. Finally, the ex vivo model also allows the testing of substances that do not cross the placental barrier, such as function-blocking antibodies. Due to these advantages, the ex vivo hindbrain culture provides an alternative and complementary method to using zebrafish embryos, which can be treated with water soluble small molecules in the aquarium water, or to in utero electroporation of embryonic brains, which requires the use of specialized equipment and is more difficult to master than the culture technique described here. Another advantage of the protocol described here is its amenability to implanting beads soaked in recombinant protein or other reagents, therefore allowing the application of a standard embryological method developed to manipulate chick embryos to a mouse model of neuronal migration. In particular, the ex vivo culture model may be applied to hindbrains of genetically engineered mice defective in specific molecules implicated in neuronal migration, such as growth or guidance factor receptors, and combined with bead implants to test if responsiveness to ligands is lost. In addition to pharmacological manipulation, the ex vivo culture protocol could also be adapted to electroporate expression vectors that could manipulate expression of genes of interest; appropriate methods for electroporation have previously been described22,23. This protocol may also be adapted to visualize neuron migration by time-lapse microscopy in hindbrain explants from transgenic mice containing fluorescently labeled FBM neurons, e.g. Isl1-Cre; Rosa26Yfp 21. Finally, this protocol may also be used to study other types of migrating neurons in the hindbrain, such as those that form the inferior olive, although this would require the use of hindbrains at older embryonic stages and may require culture for up to 48 hr, depending on neuronal viability ex vivo.

Critical steps and troubleshooting 

For the success of this protocol, it is crucial that embryos are collected early on day E11.5, closer to E11.25, when FBM neuron migration has just begun. However, it is not always possible to catch embryos at this developmental stage due to natural mating variability of mice, and accordingly, there may be some variability in the extend of FBM migration between different experiments. Variability in FBM migration may also be observed if the experiment is not completed within the time frame assigned, about 3 hr, as can be seen in Figure 3E. Hindbrain tissue from E11.25 mouse embryos is delicate. When dissecting and throughout the explant procedure, it is important to not tear the hindbrain tissue in the areas from r4-r6 where the FBM neurons are located. Due to the delicate nature of the dissection process, and because the speed at which hindbrains are placed into culture influences outcome, the procedure might take a couple of practice runs to master, in particular before precious samples or reagents are used. Finally, it is important that the hindbrain tissue is placed into an open book configuration on the culture insert, because folding of the hindbrain tissue during culture will prevent normal FBM migration (see Figure 3F).

Embryonic neurons are born in the ventricular zone of the brain, but subsequently migrate to new destinations to reach appropriate target regions that are located at a large distance. The correct positioning of neuronal cell bodies in appropriate places along the dorso-ventral and anterior-posterior axes of the developing brain is essential for the correct wiring, survival, and function of these neurons after the migratory stage1-4. Similar to the molecular mechanisms that control axon guidance5-7, combinatorial sets of attractive and repulsive cues are thought to guide migrating neurons1,8. However, due to the interactions of multiple cell types, the signals controlling neuronal migration have been less extensively studied than those involved in axon guidance, which can be studied cell autonomously. The developing hindbrain of vertebrates has been used in several recent studies to understand the molecular and cellular mechanisms of neuronal migration, for example in chick, mouse, and zebrafish1-4,9. This organ contains several different types of neurons, including several subtypes of precerebellar and motor neurons5,7,10,11.

Hindbrain motorneuron are born in the ventricular zone close to the floorplate, and they differentiate into specific subsets according to their rhombomere of origin1,12. The facial branchiomotor (FBM) neurons are generated in rhombomere (r) 4 in the hindbrain and extend their axons dorsally through an r4 exit point into the second branchial arch to innervate the facial muscles2,9,13. FBM neurons of zebrafish and mice provide excellent models to study the molecular and cellular mechanisms of neuronal migration in a process that is readily visualized, because these neurons reproducibly translocate their somata in a spatiotemporally well-defined process. In mice, FBM neurons first migrate caudally through r5 and then both caudally and ventrally to reach their final position on the pial side of the hindbrain in the territory of r6, where they form the paired nuclei of the VIIth cranial nerve (VIIn)10,11,14. In zebrafish, FBM neurons initially migrate ventrally and then change direction at the r4-r5 boundary to continue migrating towards the pial surface in a laminin-dependent manner4,12,15,16. This migration proceeds over a period of several days in development and can be divided into phases of tangential and radial migration, allowing the identification of molecules that mediate these two distinct processes. In contrast, the FBM neurons of the embryonic chick hindbrain remain in r43,13,17-19.

During their migration, FBM neurons can be identified, like other types or motor neurons, through their expression of the homoeodomain transcription factor islet 1 (ISL1)14. Thus, wholemount immunofluorescence staining or in situ hybridization for this marker at different developmental stages reveals the distinct migratory stream of FBM somata extending from r4 to r6 in the zebrafish or mouse4,15,16. Moreover, fluorescent transgenic reporters such as ISL1-GFP have been used as suitable tools to visualize migrating FBM neurons in zebrafish3,17-19. In addition to their suitability for imaging, many investigators have studied the migration of FBM neurons in developing zebrafish, because their free-living embryos can be manipulated easily with cell transplantation techniques and pharmacological compounds applied directly to the aquarium water. In contrast, the mouse embryo develops enclosed in the uterus, precluding the implantation of beads carrying guidance cues or the administration of function-blocking antibodies that do not cross the placental barrier. Moreover, pharmacological compounds administered to the pregnant mother may have undesired side effects that can indirectly impair embryogenesis. Circumventing this limitation, we have developed an ex vivo culture method for whole mouse hindbrain that is compatible with FBM neuron migration and survival for 24 hr after explanting7,16. This method allows easy pharmacological manipulation, implantation of beads carrying guidance cues or administration of function-blocking antibodies and could also be adapted to study the migration of other neuronal subtypes in the hindbrain at different developmental stages.

<table border="0" cellpadding="0" cellspacing="0" width="575">
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Eppendorf round-bottomed reagent tube, 2.0 ml (Safe-Lock)</td>
			<td style="width:111px;">VWR</td>
			<td style="width:104px;">211-2120</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Cell culture plates, 12 well</td>
			<td style="width:111px;">Thermo Scientific</td>
			<td style="width:104px;">150628</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Plastic cell culture dish, 100 mm</td>
			<td style="width:111px;">Thermo Scientific</td>
			<td style="width:104px;">150288</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">15 mm Netwell insert with Mesh Polyester Membrane</td>
			<td style="width:111px;">Corning</td>
			<td style="width:104px;">3477</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Watchmaker forceps, no. 5</td>
			<td style="width:111px;">Dumont</td>
			<td style="width:104px;">91150-20</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Phosphate buffered saline</td>
			<td style="width:111px;">Sigma</td>
			<td style="width:104px;">P4417</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Laminin mouse protein</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">23017-015</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:215px;">Primary antibody, Isl1</td>
			<td style="width:111px;">Developmental Hybridoma Bank</td>
			<td style="width:104px;">39.4D5</td>
			<td style="width:147px;">Dilution 1/100</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">AlexaFluor 488-conjugated goat anti mouse secondary antibody</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">A11029</td>
			<td style="width:147px;">Dilution 1/200</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Neurobasal medium</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">21103</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">B27 supplement (50x)&#160;</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">17504-044</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Leibovitz&#8217;s L-15</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">21083-027</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Penicillin/ streptomycin&#160;</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">15070</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Glucose</td>
			<td style="width:111px;">VWR</td>
			<td style="width:104px;">101174Y</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Heat-inactivated goat serum</td>
			<td style="width:111px;">Sigma</td>
			<td style="width:104px;">G9023</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Triton X-100</td>
			<td style="width:111px;">Sigma</td>
			<td style="width:104px;">T8787</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Paraformaldehyde</td>
			<td style="width:111px;">Sigma</td>
			<td style="width:104px;">P6148</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Slowfade Antifade Kit</td>
			<td style="width:111px;">Life Technologies</td>
			<td style="width:104px;">S-2828</td>
			<td style="width:147px;">Alternative mounting solutions may be used</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Microscope slides</td>
			<td style="width:111px;">VWR</td>
			<td style="width:104px;">631-0912</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Cover glass</td>
			<td style="width:111px;">VWR</td>
			<td style="width:104px;">631-0137</td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:215px;">Affi-Gel Heparin Beads</td>
			<td style="width:111px;">Biorad</td>
			<td style="width:104px;">153-6173</td>
			<td style="width:147px;">Glass, latex, or coloured beads may be used alternatively</td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;width:215px;">Recombinant human VEGF165</td>
			<td style="width:111px;">R&#38;D systems</td>
			<td style="width:104px;">293-VE</td>
			<td style="width:147px;">Resuspend in PBS, store aliquots at -80oC</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:215px;">Stereo Microscope, Leica MZ16</td>
			<td style="width:111px;">Leica</td>
			<td style="width:104px;"></td>
			<td style="width:147px;"></td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:215px;">Confocal laser scanning microscope&#160; LSM710</td>
			<td style="width:111px;">Zeiss</td>
			<td style="width:104px;"></td>
			<td style="width:147px;"></td>
		</tr>
	</tbody>
</table>

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.

This section illustrates examples of results that can be obtained by studying FBM neuron migration in the mouse hindbrain through ex vivo culture. We show that the FBM neurons in explanted hindbrains from day 11 mouse embryos first undergo a tangential migration (Figure 3A) and then begin to assemble the facial motor nuclei (Figure 3B), similar to their behavior in utero (see Figure 1). We further demonstrate that the implantation of a VEGF165-soaked bead attracts FBM neurons (Figures 3C and 3D), as previously shown16. Importantly, this protocol allows studying FBM migration in the absence of blood vessels or vessel-derived factors that may influence FBM migration in utero, because the nonperfused vasculature degenerates in culture16. Thus, the unspecific blood cell labelling observed when using the ISL1 mouse antibody on freshly isolated mouse hindbrains (Figure 1) is no longer present in hindbrain tissue after 24 hr in culture (Figures 3A-F). Finally, we show two examples of hindbrains that were not explanted correctly and therefore contain FBM neurons in an abnormal distribution, either because the hindbrain was not explanted soon enough after embryo isolation (Figure 3E) or because the hindbrain tissue folded up in the transwell (Figure 3F).&#160;
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51397/51397fig1highres.jpg" src="/files/ftp_upload/51397/51397fig1.jpg" /> 
Figure 1. FBM neuron migration. Confocal z-stack of wildtype mouse hindbrains after ISL1 wholemount immunolabeling and flatmounting; the hindbrain midline is indicated with an asterisk in all panels. (A) Ventricular surface of an E11.5 hindbrain in the area containing ISL1-positive FBM neurons (arrow), demonstrating their tangential migration from r4 to r6; the position of r4, r5 and r6 is indicated. (A&#8217;) Pial surface of one half of the same hindbrain in the area containing the anlage of one of the paired FBM nuclei (indicated with VIIn), as well as other ISL1-positive neuron populations. (B) Ventricular surface of an E12.5 hindbrain containing FBM neurons that are migrating tangentially (arrow); the arrowhead indicate an example of a blood vessel containing circulating cells that are unspecifically labeled by cross-reaction of the anti-mouse secondary antibody used to detect the ISL1 mouse IgG antibody. (B&#8217;) Pial surface of one half of the same E12.5 hindbrain, which contains one of the paired FBM nuclei. The midline is indicated with an asterisk in each panel. Scale bar (all panels): 200 &#956;m. V, ventral; P, pial. <a href="https://www.jove.com/files/ftp_upload/51397/51397fig1highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51397/51397fig2highres.jpg" src="/files/ftp_upload/51397/51397fig2.jpg" /> 
Figure 2. E11.5 mouse hindbrain dissection and ex vivo culture. (A-E) Key steps in the E11.5 hindbrain dissection protocol; scale bar: 1 mm. (A) Head of the embryo after it was cut away from the remainder of the embryo at forelimb level. (B) The rostral part of the head was removed and the remainder of the head tissue positioned so that the 4th ventricle (arrow) was oriented upwards. (C) The roof of the 4th ventricle was peeled away, and the hindbrain was exposed by peeling tissue beneath the hindbrain away rostrally and caudally. (D) The pial membrane was removed (note that in this example, some cervical spinal cord (SC) tissue has remained attached to the hindbrain). (E) Excess midbrain (MB) and spinal cord (SC) tissue has been removed to retain just the hindbrain. (F) Culture inserts were coated with laminin and placed into a 12 well tissue culture plate. (G) Each hindbrain was placed onto one insert and covered with media. (H) Schematic representation of the path taken by migrating FBM neurons (blue) during 24 hr of culture. <a href="https://www.jove.com/files/ftp_upload/51397/51397fig2highres.jpg" target="_blank">Click here to view larger image.</a>
<img alt="Figure 3" fo:content-width="4in" fo:src="/files/ftp_upload/51397/51397fig3highres.jpg" src="/files/ftp_upload/51397/51397fig3.jpg" /> 
Figure 3. Mouse hindbrain ex vivo culture. (A,B) An E11.5 hindbrain was cultured for 24 hr and immunofluorescently labeled for ISL1 to illustrate FBM neuron migration in an explant; both ventricular (A) and pial (B) sides of the hindbrain are shown. (C,D) E11.5 littermate hindbrains were cultured in the presence of implanted heparin bead soaked in PBS (C) or VEGF165 (D); note that FBM neurons migrated towards and onto the VEGF165 bead, and the migrating stream therefore extended further caudally compared to the untreated side of the same hindbrain or the hindbrain containing the control bead. (E,F) Examples of unsatisfactory E11.5 hindbrain explants, in which FBMs have not emigrated from r4 (E), or in which the hindbrain tissue folded during culture (F). The midline is indicated with an asterisk in each panel. Scale bar (for all panels): 200 &#956;m. V, ventral; P, pial. <a href="https://www.jove.com/files/ftp_upload/51397/51397fig3highres.jpg" target="_blank">Click here to view larger image.</a>

Watch this Scientific Journal Video about Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration at JoVE.com

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.

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 ...

mouse-hindbrain-ex-vivo-culture-to-study-facial-branchiomotor-neuron

Research

JoVE Journal

Neuroscience

11.1K Views.  University College London. 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.

Video: Mouse Hindbrain Ex Vivo Culture to Study Facial Branchiomotor Neuron Migration

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of existing mouse lines for genetic cell lineage tracing: a tamoxifen-inducible Cre line and a Cre reporter line expressing dsRed upon Cre-mediated recombination. By using a relatively low level of tamoxifen, we were able to induce recombination in a small number of cells, permitting us to follow individual cell divisions. Additionally, we observed the transcriptional response to Sonic Hedgehog (Shh) signaling using an Olig2-eGFP transgenic line 1-3 and we monitored formation of cilia by infecting the cultured slice with virus expressing the cilia marker, Sstr3-GFP 4. In order to image the neuroepithelium, we harvested embryos at E8.5, isolated the neural tube, mounted the neural slice in proper culturing conditions into the imaging chamber and performed time-lapse confocal imaging. Our ex vivo live imaging method enables us to trace single cell divisions to assess the relative timing of primary cilia formation and Shh response in a physiologically relevant manner. This method can be easily adapted using distinct fluorescent markers and provides the field the tools with which to monitor cell behavior in situ and in real time.

Ex vivo Live Imaging of Single Cell Divisions in Mouse Neuroepithelium

Here we develop the tools necessary for ex vivo live imaging to trace single cell divisions in the mouse E8.5 neuroepithelium

We developed a system that integrates live imaging of fluorescent markers and culturing slices of embryonic mouse neuroepithelium. We took advantage of ...

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.

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.

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

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 ...

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

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

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

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

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

Low-Density Primary Hippocampal Neuron Culture

The ability to probe the structure and physiology of individual nerve cells in culture is crucial to the study of neurobiology, and allows for flexibility in genetic and chemical manipulation of individual cells or defined networks. Such ease of manipulation is simpler in the reduced culture system when compared to the intact brain tissue. While many methods for the isolation and growth of these primary neurons exist, each has its own limitations. This protocol describes a method for culturing low-density and high-purity rodent embryonic hippocampal neurons on glass coverslips, which are then suspended over a monolayer of glial cells. This &#39;sandwich culture&#39; allows for exclusive long-term growth of a population of neurons while allowing for trophic support from the underlying glial monolayer. When neurons are of sufficient age or maturity level, the neuron coverslips can be flipped-out of the glial dish and used in imaging or functional assays. Neurons grown by this method typically survive for several weeks and develop extensive arbors, synaptic connections, and network properties.

This article describes the protocol for culturing low-density primary hippocampal neurons growing on glass coverslips inverted over a glial monolayer. The neuron and glial layers are separated by paraffin wax beads. The neurons grown by this method are suitable for high-resolution optical imaging and functional assays.

The ability to probe the structure and physiology of individual nerve cells in culture is crucial to the study of neurobiology, and allows for flexibility ...

The Mouse Hindbrain As a Model for Studying Embryonic Neurogenesis

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded forebrain neuroepithelium is not amenable to wholemount analysis to examine organ-wide neurogenesis patterns. Moreover, defining the mechanisms of forebrain neurogenesis is not necessarily predictive of neurogenesis in other parts of the brain; for example, due to the presence of forebrain-specific progenitor subtypes. The mouse hindbrain provides an alternative model for studying embryonic neurogenesis that is amenable to wholemount analysis, as well as tissue sections to observe the spatiotemporal distribution and behavior of neural progenitors. Moreover, it is easily dissected for other downstream applications, such as cell isolation or molecular biology analysis. As the mouse hindbrain can be readily analyzed in the vast number of cell lineage reporter and mutant mouse strains that have become available, it offers a powerful model for studying the cellular and molecular mechanisms of developmental neurogenesis in a mammalian organism. Here, we present a simple and quick method to use the mouse embryo hindbrain for analyzing mammalian neural progenitor cell (NPC) behavior in wholemount preparations and tissue sections.

This article demonstrates how the mouse embryonic hindbrain can be used as a model for studying developmental neurogenesis in both whole organ and tissue section preparations.

The mouse embryo forebrain is the most commonly employed system for studying mammalian neurogenesis during development. However, the highly folded ...

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, very few have been widely accepted. Here, we propose&#160;a new procedure&#160;adapted from the well-known ex vivo brain slice technique, which offers a closer in vivo-like scenario than in vitro preparations, for investigating the early events triggering cell degeneration, as observed in Alzheimer&#39;s disease (AD). This variation consists of simple and easily reproducible steps, which enable preservation of the anatomical cytoarchitecture of the selected brain region and its local functionality in a physiological milieu. Different anatomical areas can be obtained from the same brain, providing the opportunity to perform multiple experiments with the treatments in question in a site-, dose-, and time-dependent manner. Potential limitations&#160;which could affect the outcomes related to this methodology are related to the conservation of the tissue, i.e., the maintenance of its anatomical integrity during the slicing and incubation steps and the section thickness, which can influence the biochemical and immunohistochemical analysis. This approach can be employed for different purposes, such as exploring molecular mechanisms involved in physiological or pathological conditions, drug screening, or dose-response assays. Finally, this protocol could also reduce the number of animals employed in behavioral studies. The application reported here has been recently described and tested for the first time on ex vivo rat brain slices containing the basal forebrain (BF), which is one of the cerebral regions primarily affected in AD. Specifically, it has been demonstrated that the administration of a toxic peptide derived from the C-terminus of acetylcholinesterase (AChE) could prompt an AD-like profile, triggering, along the antero-posterior axis of the BF, a differential expression of proteins altered in AD, such as the alpha7 nicotinic receptor (&#945;7-nAChR), phosphorylated Tau (p-Tau), and amyloid beta (A&#946;).

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

We present a method which can&#160;provide further insights into the early events underlying neurodegeneration and based on the established ex-vivo brain technique, combining the advantages of in vivo and in vitro experiments. Moreover, it represents a unique opportunity for direct comparison of treated and untreated group in the same anatomical plane.

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, ...

Facial Nerve Surgery in the Rat Model to Study Axonal Inhibition and Regeneration

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial nerve can be manipulated along its entire length, from its intracranial segment to its extratemporal course. There are three primary types of nerve injury used for the experimental study of regenerative properties: nerve crush, transection, and nerve gap. The range of possible interventions is vast, including surgical manipulation of the nerve, delivery of neuroactive reagents or cells, and either central or end-organ manipulations. Advantages of this model for studying nerve regeneration include simplicity, reproducibility, interspecies consistency, reliable survival rates of the rat, and an increased anatomic size relative to murine models. Its limitations involve a more limited genetic manipulation versus the mouse model and the superlative regenerative capability of the rat, such that the facial nerve scientist must carefully assess time points for recovery and whether to translate results to higher animals and human studies. The rat model for facial nerve injury allows for functional, electrophysiological, and histomorphometric parameters for the interpretation and comparison of nerve regeneration. It thereby boasts tremendous potential toward furthering the understanding and treatment of the devastating consequences of facial nerve injury in human patients.

This protocol describes a reproducible approach to facial nerve surgery in the rat model, including descriptions of various inducible patterns of injury.

This protocol describes consistent and reproducible methods to study axonal regeneration and inhibition in a rat facial nerve injury model. The facial ...

Ex Vivo Pressurized Hippocampal Capillary-Parenchymal Arteriole Preparation for Functional Study

From subtle behavioral alterations to late-stage dementia, vascular cognitive impairment typically develops following cerebral ischemia. Stroke and cardiac arrest are remarkably sexually dimorphic diseases, and both induce cerebral ischemia. However, progress in understanding the vascular cognitive impairment, and then developing sex-specific treatments, has been partly limited by challenges in investigating the brain microcirculation from mouse models in functional studies. Here, we present an approach to examine the capillary-to-arteriole signaling in an ex vivo hippocampal capillary-parenchymal arteriole (HiCaPA) preparation from mouse brain. We describe how to isolate, cannulate, and pressurize the microcirculation to measure arteriolar diameter in response to capillary stimulation. We show which appropriate functional controls can be used to validate the HiCaPA preparation integrity and display typical results, including testing potassium as a neurovascular coupling agent and the effect of the recently characterized inhibitor of the Kir2 inward rectifying potassium channel family, ML133. Further, we compare the responses in preparations obtained from male and female mice. While these data reflect functional investigations, our approach can also be used in molecular biology, immunochemistry, and electrophysiology studies.

The present manuscript details how to isolate hippocampal arterioles and capillaries from the mouse brain and how to pressurize them for pressure myography, immunofluorescence, biochemistry, and molecular studies.

From subtle behavioral alterations to late-stage dementia, vascular cognitive impairment typically develops following cerebral ischemia. Stroke and ...

In vivo Calcium Imaging of Mouse Geniculate Ganglion Neuron Responses to Taste Stimuli

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using calcium as a proxy for neuronal activity, these techniques provide a way to monitor the responses of individual cells within large neuronal ensembles to a variety of stimuli in real time. We, and others, have applied these techniques to image the responses of individual geniculate ganglion neurons to taste stimuli applied to the tongues of live anesthetized mice. The geniculate ganglion is comprised of the cell bodies of gustatory neurons innervating the anterior tongue and palate as well as some somatosensory neurons innervating the pinna of the ear. Imaging the taste-evoked responses of individual geniculate ganglion neurons with GCaMP has provided important information about the tuning profiles of these neurons in wild-type mice as well as a way to detect peripheral taste miswiring phenotypes in genetically manipulated mice. Here we demonstrate the surgical procedure to expose the geniculate ganglion, GCaMP fluorescence image acquisition, initial steps for data analysis, and troubleshooting. This technique can be used with transgenically encoded GCaMP, or with AAV-mediated GCaMP expression, and can be modified to image particular genetic subsets of interest (i.e., Cre-mediated GCaMP expression). Overall, in vivo calcium imaging of geniculate ganglion neurons is a powerful technique for monitoring the activity of peripheral gustatory neurons and provides complementary information to more traditional whole-nerve chorda tympani recordings or taste behavior assays.

Here we present how to expose the geniculate ganglion of a live, anesthetized laboratory mouse and how to use calcium imaging to measure the responses of ensembles of these neurons to taste stimuli, allowing for multiple trials with different stimulants. This allows for in depth comparisons of which neurons respond to which tastants.

Within the last ten years, advances in genetically encoded calcium indicators (GECIs) have promoted a revolution in in vivo functional imaging. Using ...