We would like to thank the lab of A. Hadjantonakis (Sloan-Kettering Institute) and the lab of L. Niswander (U of Colorado-Denver) for helpful advice with dissection and culture techniques. This work has been supported by NRSA NS059562 to JDG and RO1NS05897 to MER in collaboration with L. Niswander and J. Nadeau (Institute for Systems Biology).

1. Preparation of culture media - all reagents are listed in Table 1
<ol>
	<li>Thaw male rat serum at 37&deg;C.</li>
	<li>Heat-Inactivate rat serum for 30 mins at 55&deg;C</li>
	<li>Centrifuge rat serum at 10K for 5 mins at room temperature</li>
	<li>Remove supernatant and mix 50:50 with phenol-red free DMEM w/ High Glucose</li>
	<li>Sterilize the media using a 0.45 um syringe filter</li>
	<li>At least 1ml of media / embryo should be prepared</li>
</ol>
2. Isolation of the uterus from the pregnant dam
<ol>
	<li>Using procedures approved by the Institutional Animal Care and Use Committee (IACUC), the pregnant dam is first anesthetized and then sacrificed by cervical dislocation</li>
	<li>Sterilize the abdomen with 70% EtOH</li>
	<li>Pinch a small path of skin along the midline, just above the nipple line, with large forceps and open the abdomen underneath your forceps with large scissors. Be careful not to damage any internal structures</li>
	<li>Use your scissors to cut laterally from the initial opening toward each side of the mouse so that the entire abdomen is open</li>
	<li>Intestines and excess fat pads can often obscure the uterus and these should be moved aside so that the top of the uterus and ovaries can be observed on each side</li>
	<li>Pinch the top of the uterus below the ovaries and use your scissors to separate the uterus from the fat, cutting from one end of the uterus to the other ovary</li>
	<li>Lift out the uterus with your forceps and place in a Petri dish with saline</li>
</ol>
3. Removal of embryos from the uterus
<ol>
	<li>Rinse the uterus with saline to remove any excess blood and trim any excess fat from the outside</li>
	<li>Transfer to a clean dish with room temp. Tyrodes saline to improve visualization</li>
	<li>Using a stereomicroscope, separate each embryo using either small scissors or if necessary fine (#5) forceps to carefully peel the uterus apart.</li>
	<li>Insert fine forceps into the space between the uterine wall and the decidua and peel back the uterine wall, being careful not to pierce the embryo</li>
</ol>
4. Removal of decidua from embryos
<ol>
	<li>Orient the decidua so that the darker portion (placenta) is facing away from you</li>
	<li>Make an incision with your forceps along the line separating the dark portion of the decidua from the lighter portion, being careful not to cut too deeply</li>
	<li>Complete the separation of the placenta from the top of the yolk sac, being careful not to tear the ectoplacental cone (EPC). At this point you should be able to visualize the Reicherts membrane on the yolk sac</li>
	<li>Continue to remove the entire decidua (lighter portion) being careful not to pierce the yolk sac</li>
	<li>Gently pinch and remove the Reicherts by peeling it away from the EPC. If the EPC is damaged or separated from the yolk sac, embryos will fail to turn normally from E8.5-E9.5</li>
	<li>With the Reicherts removed the embryo should be easily visible through the yolk sac and can be staged by counting somities.</li>
</ol>
5. Setting up the culture system
<ol>
	<li>Cut the end of a plastic transfer pipette with scissors to increase the diameter of the opening so that an embryo can be pipetted without damaging the yolk sac</li>
	<li>Fill clean roller bottles with 1 ml of media per embryo. Depending on the type of roller bottles used a maximum of 3-6 embryos can be loaded in each bottle.</li>
	<li>Transfer the embryos using your cut pipette to the media-filled roller bottle, carrying over as little Tyrodes as possible so as not to dilute the media</li>
	<li>Attach the rubber plug (bung) to the top of the glass roller bottle</li>
	<li>Insert the roller bottles into the roller culture drum so that a tight seal is formed between the plug and the drum</li>
	<li>Once all the bottles have been attached, and any empty slots in the drum have been sealed with rubber stoppers, activate the roller</li>
	<li>Open CO2 and O2 tanks and adjust them to approximately 2 psi, so that the outlet valve is releasing one bubble per second. For E8.5-E10 embryos, 20% O2 and 5% CO2 should be used.</li>
	<li>Ensure that the incubator is set to 37&deg;C and cover or close the incubator so that the embryos are shielded from light.</li>
	<li>Embryos should be checked periodically to assess their development by the addition of somities, the progression of turning, and the extent of neural tube closure</li>
	<li>After 2-3 hours in culture, pharmacological inhibitors or other treatments can be added to the media</li>
	<li>The study design should include controls, such as vehicle or inactive analogues of study reagent.</li>
</ol>
6. Evaluating development after whole embryo culture
<ol>
	<li>Turn off gas, stop roller, and remove bottles from incubator</li>
	<li>Transfer embryos back to Tyrodes or PBS in a Petri dish for evaluation under stereomicroscope</li>
	<li>The yolk sac should appear balloon-like, a heartbeat should be visible and heart rate should be &gt;120 beats per min, and circulation should be evident</li>
	<li>Photograph embryos prior to yolk sac removal if necessary</li>
	<li>Remove yolk sac from embryo by cutting a hole near the EPC and flipping the yolk sac and amnion over the head and rump of the embryo.</li>
	<li>The umbilical cord can be cut to separate the yolk sac from the embryo, and the yolk sac can be used for genotyping the embryo if necessary.</li>
	<li>Examine the embryo for somite number and neural tube defects, such as open cranial folds, incomplete closure of the face and/or an open caudal neuropore. Embryos can be staged according to the Theiler classification of multiple morphogenetic changes (<a href="http://genex.hgu.mrc.ac.uk/Databases/Anatomy/MAstaging.html">http://www.emouseatlas.org/ emap/home.html</a>) Again, a photographic record of embryo development is useful.</li>
</ol>
7. Notes
<ol>
	<li>A high quality dissection is essential, because any nick or tears in the yolk sac will prevent successful turning and inhibit normal development. The EPC can also be damaged during removal of the Reichert's membrane (RM), and if the EPC is even partially separated from the yolk sac, embryos will not develop normally. However, incomplete removal of the RM can cause embryos to stick together and/or RM cells may overgrow in culture and constrict yolk sac expansion, both of which would prevent normal development.</li>
	<li>Clean and sharp forceps are the secret to getting a good dissection, because dull or bent forceps make the procedure significantly more difficult. Protein deposits on instruments will lead to tissue damage.</li>
	<li>Clean, sterile glass bottles are also essential to prevent embryos from sticking to the side wall and being damaged. Prior to starting the experiment, bottles should be scrubbed with soap and water, rinsed in dH2O, and microwaved with a small amount of water in each bottle until the water boils. Bottles are then rinsed with 70% EtOH and allowed to air dry.</li>
	<li>Work as quickly as possible, because the sooner the embryo can be transferred to the culture media and warmed back to 37&deg;C the better survival.</li>
	<li>Complete dissecting the entire litter before transferring any embryos to roller bottles so that each embryo is at room temperature for the same amount of time to avoid creating any discrepancies in embryo growth</li>
	<li>Maintain a sterile environment to prevent bacterial contamination of the cultures. Our culture experiments have been performed without antibiotic, but many labs add antibiotic to their media to prevent bacterial growth. The medium should appear as clear at the end of the experiment as it was at the beginning of the experiment. If media has become cloudy or there is a visible precipitate, there is likely a contamination that has infected your culture.</li>
	<li>Ineffective gas exchange can also delay embryos development, so be sure to have a reliable regulator that will ensure continuous delivery.</li>
	<li>Embryos develop approximately 50% slower in ex vivo culture, therefore it is essential to count somites to ensure that you have reached the appropriate stage of development for completion of the experiment.</li>
</ol>
8. Representative results:
The appearance of embryos pre- and post- roller culture is illustrated in Figure 1. At the time of dissection, embryos should be in an unturned configuration (Fig. 1A,D) where the tail is behind the head folds. After 36 hrs in culture, embryos should have completed turning so that they are in the C-curved, fetal position, where the tail is in front of the head (Fig 1B,C,E,F). Pharmacological manipulation with RhoA kinase inhibitor (Y-27632), a known inhibitor of convergent extension during neurulation (Ybot-Gonzalez, 2007), results in a shortening of the embryos along their rostral-caudal axis (Fig. 1E,F) and inhibits cranial neural fold closure. Our data show that increasing doses of Y-27632 progressively impairs cranial fold closure (Figure 2A) and shortens the body axis (Fig. 2B), consistent with the role of downstream RhoA signaling in cranial neurulation and convergent extension.
<img alt="Figure 1" src="/files/ftp_upload/3132/3132fig1.jpg" /> 
Figure 1. Appearance of cultures embryos and manipulation with the pharmacological inhibitor Y-27632. (A,D) Dissected embryos at E8.5 prior to whole embryo culture (B,E) Embryos at E10 with the yolk sac still intact, subsequent to 36hrs of roller culture. (C,F) The yolk sac has been removed to illustrate successful turning and neural tube closure. Embryos that were treated with the Rho kinase inhibitor Y-27632 (E,F) failed to undergo proper convergent extension and illustrate a shortened body axis.
<img alt="Figure 2" src="/files/ftp_upload/3132/3132fig2.jpg" /> 
Figure 2. Effect of RhoA kinase inhibitor on cranial neural tube closure and axis elongation. (A) The percentage of embryos that were able to successfully close their cranial folds (%NTC= percentage neural tube closure) is compared to the dose of Y-27632 added to culture media. (B) The distance between the otic vesicle and forelimb was significantly reduced at increasing doses of Y-27632 (p&lt;.05).

<ol>
	<li>Cockroft, D. L., Copp, A. J., Cockroft, D. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissection+and+culture+of+post+implantation+embryos.">Dissection and culture of post implantation embryos.</a> Mammalian Postimplantation Embryos A Practical Approach. , (1990).</li><li>Gray, J. D., Ross, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanistic+insights+into+folate+supplementation+from+Crooked+tail+and+other+NTD-prone+mutant+mice.">Mechanistic insights into folate supplementation from Crooked tail and other NTD-prone mutant mice.</a> Birth Defects Res. A. Clin. Mol. Teratol. 85, 314-321 (2009).</li><li>Kaufman, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Atlas of Mouse Development. , (1992).</li><li>Ybot-Gonzalez, P., Savery, D., Gerrelli, D., Signore, M., Mitchell, C. E., Faux, C. H., Greene, N. D., Copp, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convergent+extension,+planar-cell-polarity+signalling+and+initiation+of+mouse+neural+tube+closure.">Convergent extension, planar-cell-polarity signalling and initiation of mouse neural tube closure.</a> Development. 134, 789-799 (2007).</li><li>Ross, M. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gene-environment+interactions,+folate+metabolism+and+the+embryonic+nervous+system.">Gene-environment interactions, folate metabolism and the embryonic nervous system.</a> Wiley Interdiscip Rev Syst Biol Med. 2, 471-480 (2010).</li></ol>

No conflicts of interest declared.

The ability to separate maternal, intrauterine factors from those that are intrinsic to the embryo during its development is an important tool for studying all stages of embryogenesis. Here we have analyzed the effects of a small molecule inhibitor of RhoA kinase on cranial neurulation ex utero, thereby removing the variable of maternal metabolism of the drug. This pharmacological manipulation has a profound effect on cranial neurulation and convergent extension. Sensitivity to this compound can be compared among different genetic mouse mutants. The method presented here can also be applied to studies of other molecular pathways in development, allowing the direct manipulation of cellular function in embryos using a variety of reagents.

<table border="1">
	<tbody>
		<tr>
			<th>Name</th>
			<th>Company</th>
			<th>Catalogue number</th>
			<th>Comments</th>
		</tr>
		<tr>
			<td>Stereomicroscope for Dissection</td>
			<td>Leica</td>
			<td>M165</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Precision Incubator	</td>
			<td>BTC Engineering (U.K.)</td>
			<td>BTC 001</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rotating Bottle Culture Unit</td>
			<td>BTC</td>
			<td>BTC 003</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glass Bottles for Rotating Unit</td>
			<td>BTC</td>
			<td>BTC 005</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Silicone Rubber Bung</td>
			<td>BTC</td>
			<td>BTC 007</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Silicone Rubber Cork</td>
			<td>BTC</td>
			<td>BTC 008</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gas Bubbler Inlet</td>
			<td>BTC</td>
			<td>BTC 012</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gas Bubbler Inlet Trap</td>
			<td>BTC</td>
			<td>BTC 013</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gas Bubbler Outlet</td>
			<td>BTC</td>
			<td>BTC 014</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gas Cylinder</td>
			<td>TechAir</td>
			<td>&nbsp;</td>
			<td>20% O2, 5% CO2</td>
		</tr>
		<tr>
			<td>Gas Regulator</td>
			<td>TechAir</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Male Rat Serum</td>
			<td>Harlan Labs</td>
			<td>4520</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>DMEM w/o phenol red</td>
			<td>Invitrogen</td>
			<td>31053</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>10 mL Syringe</td>
			<td>BD</td>
			<td>309604</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Syringe Filter</td>
			<td>Nalgene</td>
			<td>190-2545</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Large Dissecting Scissors (Blunt tip)</td>
			<td>VWR</td>
			<td>82027-594</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Extra Fine Bonn Scissors</td>
			<td>Fine Scientific Tools (FST)</td>
			<td>14084-08</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dumot #5 forceps</td>
			<td>FST</td>
			<td>11252-50</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tyrode's Saline</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>As described in Cockroft 1990</td>
		</tr>
		<tr>
			<td>10 cm Disposable Petri Dish</td>
			<td>BD</td>
			<td>351029	</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

neural tube closure in mouse whole embryo culture

Genetic mouse models are an important tool in the study of mammalian neural tube closure (Gray &amp; Ross, 2009; Ross, 2010). However, the study of mouse embryos in utero is limited by our inability to directly pharmacologically manipulate the embryos in isolation from the effects of maternal metabolism on the reagent of interest. Whether using a small molecule, recombinant protein, or siRNA, delivery of these substances to the mother, through the diet or by injection will subject these unstable compounds to a variety of bodily defenses that could prevent them from reaching the embryo. Investigations in cultures of whole embryos can be used to separate maternal from intrinsic fetal effects on development.
Here, we present a method for culturing mouse embryos using highly enriched media in a roller incubator apparatus that allows for normal neural tube closure after dissection (Crockett, 1990). Once in culture, embryos can be manipulated using conventional in vitro techniques that would not otherwise be possible if the embryos were still in utero. Embryo siblings can be collected at various time points to study different aspects of neurulation, occurring from E7-7.5 (neural plate formation, just prior to the initiation of neurulation) to E9.5-10 (at the conclusion of cranial fold and caudal neuropore closure, Kaufman, 1992). In this protocol, we demonstrate our method for dissecting embryos at timepoints that are optimal for the study of cranial neurulation. Embryos will be dissected at E8.5 (approx. 10-12 somities), after the initiation of neural tube closure but prior to embryo turning and cranial neural fold closure, and maintained in culture till E10 (26-28 somities), when cranial neurulation should be complete.

Watch this Scientific Journal Video about Neural Tube Closure in Mouse Whole Embryo Culture at JoVE.com

Neural Tube Closure in Mouse Whole Embryo Culture

A method allowing for direct pharmacological manipulation of mouse embryos during neurulation that bypasses maternal metabolism is described. The technique can be adapted to study different aspects of neurulation by varying the time point and pharmacological agent.

Genetic mouse models are an important tool in the study of mammalian neural tube closure (Gray &amp; Ross, 2009; Ross, 2010).  However, the study of mouse ...

neural-tube-closure-in-mouse-whole-embryo-culture

Research

JoVE Journal

Neuroscience

21.4K vues.  Weill Cornell Medical College. Une méthode permettant la manipulation pharmacologique directe d'embryons de souris au cours neurulation qui contourne le métabolisme de la mère est décrite. La technique peut être adaptée pour étudier les différents aspects de la neurulation en faisant varier le point dans le temps et l'agent pharmacologique.

Vidéo: Neural Tube Closure in Mouse Whole Embryo Culture

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Production of Chick Embryo Extract for the Cultivation of Murine Neural Crest Stem Cells

The neural crest arises from the neuro-ectoderm during embryogenesis and persists only temporarily. Early experiments already proofed pluripotent progenitor cells to be an integral part of the neural crest1. Phenotypically, neural crest stem cells (NCSC) are defined by simultaneously expressing p75 (low-affine nerve growth factor receptor, LNGFR) and SOX10 during their migration from the neural crest2,3,4,5. These progenitor cells can differentiate into smooth muscle cells, chromaffin cells, neurons and glial cells, as well as melanocytes, cartilage and bone6,7,8,9. To cultivate NCSC in vitro, a special neural crest stem cell medium (NCSCM) is required10. The most complex part of the NCSCM is the preparation of chick embryo extract (CEE) representing an essential source of growth factors for the NCSC as well as for other types of neural explants. Other NCSCM ingredients beside CEE are commercially available. Producing CCE using laboratory standard equipment it is of high importance to know about the challenging details as the isolation, maceration, centrifugation, and filtration processes. In this protocol we describe accurate techniques to produce a maximized amount of pure and high quality CEE.

To cultivate neural crest stem cells (NCSC) in vitro, a special medium (NCSCM) is required. Essential part of NCSCM is chick embryo extract (CEE). We here describe accurate techniques to produce a maximized amount of pure and high quality CEE, including details as the isolation, maceration, centrifugation, and filtration processes.

The neural crest arises from the neuro-ectoderm during embryogenesis and persists only temporarily. Early experiments already proofed pluripotent ...

Establishing Embryonic Mouse Neural Stem Cell Culture Using the Neurosphere Assay

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life span of the animal. They can potentially replace cells that are lost in the aging process or in the process of injury and disease. The existence of neural stem cells (NSCs) was first described by Reynolds and Weiss (1992) in the adult mammalian central nervous system (CNS) using a novel serum&#8208;free culture system, the neurosphere assay (NSA). Using this assay, it is also feasible to isolate and expand NSCs from different regions of the embryonic CNS. These in vitro expanded NSCs are multipotent and can give rise to the three major cell types of the CNS. While the NSA seems relatively simple to perform, attention to the procedures demonstrated here is required in order to achieve reliable and consistent results. This video practically demonstrates NSA to generate and expand NSCs from embryonic day 14-mouse brain tissue and provides technical details so one can achieve reproducible neurosphere cultures. The procedure includes harvesting E14 mouse embryos, brain microdissection to harvest the ganglionic eminences, dissociation of the harvested tissue in NSC medium to gain a single cell suspension, and finally plating cells in NSA culture. After 5-7 days in culture, the resulting primary neurospheres are passaged to further expand the number of the NSCs for future experiments.

This video protocol demonstrates the application of the neurosphere assay for the isolation and expansion of neural stem cells from the ganglionic eminences of embryonic day 14-mouse brain.

In mammalians, stem cells acts as a source of undifferentiated cells to maintain cell genesis and renewal in different tissues and organs during the life ...

Localizing Protein in 3D Neural Stem Cell Culture: a Hybrid Visualization Methodology

The importance of 3-dimensional (3D) topography in influencing neural stem and progenitor cell (NPC) phenotype is widely acknowledged yet challenging to study. When dissociated from embryonic or post-natal brain, single NPCs will proliferate in suspension to form neurospheres. Daughter cells within these cultures spontaneously adopt distinct developmental lineages (neurons, oligodendrocytes, and astrocytes) over the course of expansion despite being exposed to the same extracellular milieu. This progression recapitulates many of the stages observed over the course of neurogenesis and gliogenesis in post-natal brain and is often used to study basic NPC biology within a controlled environment. Assessing the full impact of 3D topography and cellular positioning within these cultures on NPC fate is, however, difficult. To localize target proteins and identify NPC lineages by immunocytochemistry, free-floating neurospheres must be plated on a substrate or serially sectioned. This processing is required to ensure equivalent cell permeabilization and antibody access throughout the sphere. As a result, 2D epifluorescent images of cryosections or confocal reconstructions of 3D Z-stacks can only provide spatial information about cell position within discrete physical or digital 3D slices and do not visualize cellular position in the intact sphere. Here, to reiterate the topography of the neurosphere culture and permit spatial analysis of protein expression throughout the entire culture, we present a protocol for isolation, expansion, and serial sectioning of post-natal hippocampal neurospheres suitable for epifluorescent or confocal immunodetection of target proteins. Connexin29 (Cx29) is analyzed as an example. Next, using a hybrid of graphic editing and 3D modelling softwares rigorously applied to maintain biological detail, we describe how to re-assemble the 3D structural positioning of these images and digitally map labelled cells within the complete neurosphere. This methodology enables visualization and analysis of the cellular position of target proteins and cells throughout the entire 3D culture topography and will facilitate a more detailed analysis of the spatial relationships between cells over the course of neurogenesis and gliogenesis in vitro.
Both Imbeault and Valenzuela contributed equally and should be considered joint first authors.

Here, we describe how to produce, expand, and immunolabel postnatal hippocampal neural progenitor cells (NPCs) in three-dimensional (3D) culture. Next, using hybrid visualization technologies, we demonstrate how digital images of immunolabelled cryosections can be used to reconstruct and map the spatial position of immunopositive cells throughout the entire 3D neurosphere.

The importance of 3-dimensional (3D) topography in influencing neural stem and progenitor cell (NPC) phenotype is widely acknowledged yet challenging to ...

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Isolation and Culture of Neural Crest Cells from Embryonic Murine Neural Tube

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to mesenchymal transition (EMT) and migrates throughout the embryo, giving rise to diverse cell types 1-3. NC also has the unique ability to influence the differentiation and maturation of target organs4-6. When explanted in vitro, NC progenitors undergo self-renewal, migrate and differentiate into a variety of tissue types including neurons, glia, smooth muscle cells, cartilage and bone. 
NC multipotency was first described from explants of the avian neural tube7-9. In vitro isolation of NC cells facilitates the study of NC dynamics including proliferation, migration, and multipotency. Further work in the avian and rat systems demonstrated that explanted NC cells retain their NC potential when transplanted back into the embryo10-13. Because these inherent cellular properties are preserved in explanted NC progenitors, the neural tube explant assay provides an attractive option for studying the NC in vitro. 
To attain a better understanding of the mammalian NC, many methods have been employed to isolate NC populations. NC-derived progenitors can be cultured from post-migratory locations in both the embryo and adult to study the dynamics of post-migratory NC progenitors11,14-20, however isolation of NC progenitors as they emigrate from the neural tube provides optimal preservation of NC cell potential and migratory properties13,21,22. Some protocols employ fluorescence activated cell sorting (FACS) to isolate a NC population enriched for particular progenitors11,13,14,17. However, when starting with early stage embryos, cell numbers adequate for analyses are difficult to obtain with FACS, complicating the isolation of early NC populations from individual embryos. Here, we describe an approach that does not rely on FACS and results in an approximately 96% pure NC population based on a Wnt1-Cre activated lineage reporter23. 
The method presented here is adapted from protocols optimized for the culture of rat NC11,13. The advantages of this protocol compared to previous methods are that 1) the cells are not grown on a feeder layer, 2) FACS is not required to obtain a relatively pure NC population, 3) premigratory NC cells are isolated and 4) results are easily quantified. Furthermore, this protocol can be used for isolation of NC from any mutant mouse model, facilitating the study of NC characteristics with different genetic manipulations. The limitation of this approach is that the NC is removed from the context of the embryo, which is known to influence the survival, migration and differentiation of the NC2,24-28.

Isolation of embryonic neural crest from the neural tube facilitates the use of in vitro methods for studying migration, self-renewal, and multipotency of neural crest.

The embryonic neural crest (NC) is a multipotent progenitor population that originates at the dorsal aspect of the neural tube, undergoes an epithelial to ...

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are located in the dorsal spinal cord and send their axons along stereotyped trajectories. Commissural axons initially project ventrally towards and then across the floorplate. After crossing the midline, these axons make a sharp rostral turn and project longitudinally towards the brain. Each of these steps is regulated by the coordinated activities of attractive and repulsive guidance cues. The correct interpretation of these cues is crucial to the guidance of axons along their demarcated pathway. Thus, the physiological contribution of a particular molecule to commissural axon guidance is ideally investigated in the context of the living embryo. Accordingly, gene knockdown in vivo must be precisely controlled in order to carefully distinguish axon guidance activities of genes that may play multiple roles during development.
Here, we describe a method to knockdown gene expression in the chicken neural tube in a cell type-specific, traceable manner. We use novel plasmid vectors3 harboring cell type-specific promoters/enhancers that drive the expression of a fluorescent protein marker, followed directly by a miR30-RNAi transcript4 (located within the 3'-UTR of the cDNA encoding the fluorescent protein) (Figure 1). When electroporated into the developing neural tube, these vectors elicit efficient downregulation of gene expression and express bright fluorescent marker proteins to enable direct tracing of the cells experiencing knockdown3. Mixing different RNAi vectors prior to electroporation allows the simultaneous knockdown of two or more genes in independent regions of the spinal cord. This permits complex cellular and molecular interactions to be examined during development, in a manner that is fast, simple, precise and inexpensive. In combination with DiI tracing of commissural axon trajectories in open-book preparations5, this method is a useful tool for in vivo studies of the cellular and molecular mechanisms of commissural axon growth and guidance. In principle, any promoter/enhancer could be used, potentially making the technique more widely applicable for in vivo studies of gene function during development6.
This video first demonstrates how to handle and window eggs, the injection of DNA plasmids into the neural tube and the electroporation procedure. To investigate commissural axon guidance, the spinal cord is removed from the embryo as an open-book preparation, fixed, and injected with DiI to enable axon pathways to be traced. The spinal cord is mounted between coverslips and visualized using confocal microscopy.

In ovo Electroporation of miRNA-based Plasmids in the Developing Neural Tube and Assessment of Phenotypes by DiI Injection in Open-book Preparations

A method by which gene expression in the neural tube can be downregulated in a cell type-specific, traceable manner is described. We demonstrate how in ovo electroporation of microRNA-based plasmids that elicit spatiotemporally controlled RNA interference can be used to investigate commissural axon guidance in the developing neural tube.

Commissural dI1 neurons have been extensively studied to elucidate the mechanisms underlying axon guidance during development1,2. These neurons are ...

Chicken Embryo Spinal Cord Slice Culture Protocol

Slice cultures can facilitate the manipulation of embryo development both pharmacologically and through gene manipulations. In this reduced system, potential lethal side effects due to systemic drug applications can be overcome. However, culture conditions must ensure that normal development proceeds within the reduced environment of the slice. We have focused on the development of the spinal cord, particularly that of spinal motor neurons. We systematically varied culture conditions of chicken embryo slices from the point at which most spinal motor neurons had been born. We assayed the number and type of motor neurons that survived during the culture period and the position of those motor neurons compared to that in vivo. We found that serum type and neurotrophic factors were required during the culture period and were able to keep motor neurons alive for at least 24 hr and allow those motor neurons to migrate to appropriate positions in the spinal cord. We present these culture conditions and the methodology of preparing the embryo slice cultures using eviscerated chicken embryos embedded in agarose and sliced using a vibratome.

Slice cultures facilitate the manipulation of embryo development by gene and pharmacological perturbations. However, culture conditions must ensure that normal development can proceed within the reduced environment of the slice. We illustrate a protocol that facilitates normal spinal cord development to proceed for at least 24 hr.

Slice cultures can facilitate the manipulation of  embryo development both pharmacologically and through gene manipulations. In  this reduced system, ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...